【#1】Kỹ Thuật Trồng Cây Đu Đủ Cảnh Trong Chậu

Đúng như tên gọi ‘đu đủ’, cái tên thể hiện một cuộc sống đủ đầy bền vững, không thừa cũng chẳng thiếu, chính vì vậy mấy năm gần đây thú chơi đu đủ cảnh lên ngôi trong dịp Tết. Để có một chậu đu đủ cảnh được coi là đẹp với đủ hoa, lá và quả chơi Tết, người làm vườn cần có kỹ thuật trồng cây và một số biện pháp chăm sóc cơ bản:

Chọn đúng thời vụ gieo trồng

Từ trồng bầu cây lên chậu đến khi chậu cây có giá trị sử dụng làm cảnh cần khoảng thời gian từ 7-8 tháng. Nếu trồng cho mục đích kinh doanh, cần tính toán thời vụ trồng để có chậu cây cung ứng kịp thời cho nhu cầu thị trường trong dịp tết Nguyên Đán, khi đó giá trị cảnh của đu đủ sẽ cao gấp 20-30 lần so với trồng đủ đủ thuần trên cùng diện tích đất để thu quả bán ăn tươi. Ở các tỉnh đồng bằng sông Hồng thời vụ trồng đu đủ cảnh tốt nhất bắt đầu từ cuối tháng 4, đầu tháng 5 Âm lịch để kịp thời thu hoạch vào dịp Tết.

Chọn giống cây trồng hợp lý

Kỹ thuật trồng cây và chọn giống cây trồng

Ngoài việc cần gieo trồng đúng thời vụ, thì yếu tố giống cây trồng rất quan trọng. Chọn giống đu đủ lai F1, loại chuyên trồng làm cảnh: Cây lùn, lóng đốt ngắn, giống sinh trưởng khỏe, khả năng chống chịu tốt với điều kiện bất thuận, sai hoa, nhiều quả, chất lượng quả cao, trồng cây nào có quả cây đó (thường là các giống có nguồn gốc từ Thái Lan hoặc Đài Loan).

Chuẩn bị chậu và đất để trồng cây

Về chậu trồng cây, lựa chọn các chậu cây được làm bằng sứ hoặc các chậu xi măng chuyên dụng cho cây cảnh. Yêu cầu kích thước cho mỗi chậu để trồng cây là 90 x 40 x 40cm. Ngoài ra, ở mỗi chậu phải có các lỗ thoát nước ở cuối đáy và xung quanh.

Đất trồng, gồm hỗn hợp đất thịt ải, xỉ than tỷ lệ 3:1, ủ kỹ 12-15 ngày trước khi đưa vào chậu, 1m3 hỗn hợp này đủ cho 11-12 chậu đu đủ cảnh. Yêu cầu, đất thịt ải phải là đất mới chưa qua gieo trồng bất cứ loại rau màu nào. Đây là khâu kỹ thuật then chốt, có ý nghĩa quyết định tỷ lệ sống của cây đu đủ trên chậu sau trồng.

Ươm giâm bầu cây và trồng cây con lên chậu

Kỹ thuật trồng cây và ươm giâm bầu cây

Trồng đu đủ cảnh nhất thiết phải trồng bằng cây con đã gieo ươm trong bầu, có thể mua giống cây đã gieo ươm trong bầu, loại bầu có kích thước (10-12cm) x (12-15cm), cây giống to mập, khỏe, sạch bệnh có từ 4-5 cặp lá, cao 10-15cm, sau đó đưa về giâm lại trong vườn nhà, nơi cao ráo, thoáng mát.

Hàng ngày tưới nhẹ, bón thúc 1 lần sau giâm cây 2-3 ngày bằng Super lân, liều lượng 100-150g Super lân pha cho bình 8-12 lít nước, phun ướt đều cho các bầu cây, nên kết hợp phun cùng một số thuốc bảo vệ thực vật phòng ngừa nhện đỏ, rệp sáp và bệnh khảm. Khoảng 12-15 ngày sau giâm có thể đưa cây trồng lên chậu.

Trồng cây lên chậu, đổ hỗn hợp đất, xỉ than đã chuẩn bị trước vào chậu và cách miệng 5-7cm. Bón lót mỗi chậu 0,5kg NPK (15-9-17+TE), và 0,2kg vôi bột (trộn đều trong đất trước khi đưa vào chậu). Nhấc nhẹ bầu cây, dùng dao sắc rạch nhẹ vừa đứt lớp vỏ bao bầu một đường từ trên xuống sát đáy bấu, để định hướng sự phát triển rễ cây trong chậu sau này.

Đặt bầu cây (còn nguyên túi nilon bao ngoài) trồng ngay ngắn trong chậu, phủ đất kín bầu cây, nén chặt nhẹ, tưới dưỡng ẩm hàng ngày. Đặt các chậu cây trong vườn so le cách nhau 2m x (1,5-1,7m). Trong quá trình sinh trưởng, bộ rễ cây sẽ bị hạn chế phát triển bởi phần túi nilon còn bao bầu, chủ yếu phát triển tập trung theo hướng mở của vỏ nilon bao bầu rạch trước đó.

Biện pháp chăm sóc và uốn vít cây

Kỹ thuật trồng cây và uốn vít cây

Sau khi trồng cây lên chậu cần tưới ẩm ngày 1 – 2 lần, dùng rơm, rạ che phủ mặt chậu để hạn chế bốc thoát hơi nước và giữ ẩm cho cây. Định kỳ 15 ngày bón thúc phân hữu cơ, vô cơ và 30 ngày bón vôi bột, rắc trực tiếp lên mặt chậu xa gốc cây. Liều lượng mỗi lần bón cho 1 gốc: 0,2kg vôi bột, 0,2kg NPK Đầu Trâu (15-9-17+TE) + 1-1,5 phân hữu cơ hoặc 0,5kg ngô, đậu tương, giai đoạn cây mang quả bón tăng lượng NPK lên 0,3kg.

Uốn vít cây, từ 25 – 30 ngày sau trồng, tiến hành uốn vít cây. Dùng dây mềm, chắc, to bản (2 – 3cm), không co dãn, buộc vít cây tại vị trí 3/4 thân (từ gốc), kéo ngả dần về hướng đã định, ghim cố định dây chặt xuống đất, cần uốn vít dần dần để cây không bị gãy. Sau khoảng 3 tháng thì cây sẽ nghiêng 30 – 35 độ so với mặt đất.

Chúc bà con nông dân, làm vườn có một mùa bội thu!

Tâm Đinh

Nên đọc

【#2】Ngọc Lan: “kiều Nữ” Mê Trai Đẹp!

Là một trong những diễn viên trẻ được yêu thích nhất hiện nay qua hàng loạt những vai diễn trong các bộ phim truyền hình cũng như trên sân khấu kịch, Ngọc Lan được khán giả ưu ái gọi tên “kiều nữ” – vai diễn để lại ấn tượng nhất trong sự nghiệp của cô tính đến nay. Chia sẻ với 2Idol Ngọc Lan cho biết, cô thấy mình thực sự may mắn khi được góp mặt trong bộ phim “Kiều nữ và đại gia”, dù sau này có thêm nhiều vai Hồng (Âm tính, Cổng mặt trời) nhưng Hồng “kiều nữ” vẫn là bước ngoặt đáng nhớ với cô.

Để có được vị trí như ngày hôm nay, Ngọc Lan cũng phải trải qua hàng loạt những “sóng gió” trong nghề. Cô cho biết từ một hình bìa báo trong lễ giỗ Tổ Hùng Vương mà cô vinh dự cùng vài người bạn lên dâng hương, mà một đạo diễn đã để ý và mở đường cho Ngọc Lan tham gia nghệ thuật. Bắt đầu bằng công việc làm người mẫu và đóng clip ca nhạc, cho tới năm 18 tuổi thì Ngọc Lan có được một vai phụ trong phim “Công ty thời trang”. Kể từ đó, những vai diễn liên tiếp đến với cô và cho tới nay gia tài phim ảnh của Ngọc Lan cũng kha khá.

Là một người sống tình cảm, Ngọc Lan luôn luôn hướng mình về với gia đình. Mất ba từ nhỏ, một mình mẹ chắt chiu nuôi cô khôn lớn. Vì vậy mẹ chính là người mà Ngọc Lan trân trọng và yêu thương nhất. Tính cần cù, chăm chỉ, tiết kiệm của cô cũng ảnh hưởng từ mẹ. Chính vì thế, catse đóng phim, đi diễn cô đều dành dụm, tích góp để mua nhà. Cô tự hào vì hiện tại đã có một căn hộ khang trang, ấm cúng ở quận 7 – kết quả của những năm miệt mài trên phim trường với hàng chục vai diễn .

Chia sẻ với 2Idol, Ngọc Lan cũng thẳng thắn cho biết mình có được như ngày hôm nay là do bản thân cô đã phải quyết tâm vượt qua những lời chê bai, dè bỉu hồi mới chập chững vào nghề. Đó cũng chính là Ngọc Lan luôn nỗ lực, cố gắng trưởng thành hơn qua từng vai diễn dù phim truyền hình hay trên sân khấu kịch. Cô cũng bày tỏ mong muốn được sống mãi với nghề và có những vai diễn để đời. “Còn xinh đẹp là còn muốn đóng phim” – Ngọc Lan vui vẻ tâm sự.

Là một người sống tình cảm nên “kiều nữ” cũng khá lận đận trong tình duyên. Ngọc Lan đã trải qua không dưới ba mối tình và đều để lại cho cô những kỷ niệm khó quên. Cô hồ hởi bật mí mẫu người đàn ông lý tưởng của mình phải là những chàng trai đẹp, độc, vui vẻ và biết cảm thông! Là một diễn viên, lịch làm việc của cô kín mít trên phim trường từ ngày này qua tháng khác nên chắc hẳn sẽ rất cần một người biết chia sẻ, lắng nghe và cảm thông.

Không chỉ thẳng thắn trả lời hết những thắc mắc của khán giả, Ngọc Lan còn khiến phim trường 2 Idol tưng bừng hẳn lên nhờ khiếu hài hước và những màn tung hứng ăn ý cùng Khởi My, Hoàng Phi. Lần đầu tiên khán giả sẽ bất ngờ khi thấy Ngọc Lan hát cải lương Lan và Điệp cực mùi mẫn bằng cả giọng Nam và Bắc. Chưa hết, Ngọc Lan còn khiến fan “tâm phục, khẩu phục” khi hóa thân xuất sắc vào những vai diễn “kiều nữ” khác nhau : “kiều nữ” ham tiền, “kiều nữ” mê trai đẹp, “kiều nữ” lẳng lơ, “kiều nữ” thất tình …

Chương trình 2Idol với sự góp mặt của Ngọc Lan sẽ phát sóng lúc 20h50, trên VTV9 tối chủ nhật, ngày 18/8.

【#3】Trồng Lan Mokara Cắt Cành Theo Công Nghệ Mới: Mô Hình Hiệu Quả

(Tin Môi Trường) – Anh Lê Văn Đạt (khu phố 2, phường Định Hòa, TP.Thủ Dầu Một, Bình Dương) là 1 trong 5 hội viên tiêu biểu của Câu lạc bộ (CLB) Trang trại hoa lan Bình Dương được chọn tham gia dự án “Trồng lan Mokara cắt cành theo công nghệ mới” và làm mô hình trình diễn. Vườn lan của anh Đạt có diện tích 1.000m2, trồng 7.000 cây lan Mokara với các giống được thị trường ưa chuộng như vàng chanh, đỏ quặp, vàng nến…

Với hệ thống tưới phun sương bán tự động và tuân thủ chặt chẽ quy trình kỹ thuật từ chọn giống, bón phân cho đến phòng trừ sâu bệnh nên vườn lan của anh đã sinh trưởng và phát triển rất tốt. So với cách trồng bình thường, cây lan trong dự án của anh phát triển được 2 – 4 lá/tháng (bình thường 1- 2 lá/tháng), lá có màu xanh bóng đẹp, bản lá to bề ngang và dài hơn. Số cành hoa cũng phát triển nhiều hơn bình thường từ 2 – 6 cành tùy theo giống lan, sâu bệnh và cỏ dại được kiểm soát chặt chẽ, không bị gây hại. Hiện nay vườn lan của anh cho thu hoạch khoảng 1.000 cành/tháng với số tiền thu được gần 10 triệu đồng.

Anh Đạt cho biết, một trong những khó khăn lớn nhất của người trồng lan, nhất là với cây lan Mokara là vốn đầu tư ban đầu tương đối cao, mỗi cây lan giống từ 50.000 – 100.000 đồng/ cây. Theo dự toán của CLB Trang trại hoa lan Bình Dương, để trồng 100m2 cây lan Mokara phải tốn khoảng 70 triệu đồng và sau 3 năm mới có thể thu hồi được vốn. Nhằm nghiên cứu xây dựng quy trình trồng lan có tính khoa học và thiết thực, CLB đã tiến hành thực hiện đề tài “Trồng lan Mokara cắt cành theo công nghệ mới” (thực hiện từ năm 2011), cho 5 hộ vay vốn ưu đãi của Quỹ Phát triển khoa học – công nghệ tỉnh.

Đến nay đã giải ngân 1,5 tỷ đồng cho 5 hộ vay với lãi suất 5%/ năm (lãi trả 1 quý 1 lần). Các hội viên thực hiện dự án đã trả lãi hai năm ( 2011-2013) là 142.128.000 đồng và số tiền gốc là 600 triệu đồng, số còn lại phải trả vào năm 2014-2015 là 900 triệu đồng cùng với số lãi hàng quý. Với sự sinh trưởng phát triển tốt của các vườn lan điểm của dự án thì việc trả vốn gốc và lãi hàng quý được bảo đảm. Hiện 5 mô hình điểm “Trồng lan Mokara cắt cành theo công nghệ mới” của CLB có diện tích 3.000m2, trồng 21.000 cây lan Mokara. Đến nay, các mô hình này được đánh giá là sinh trưởng phát triển tốt, đang ở giai đoạn kinh doanh, hiệu quả rất tốt. Các mô hình điểm này đã có ảnh hưởng tốt và lan tỏa ra các hộ trồng lan trong CLB.

Ý kiến bạn đọc về: Trồng lan mokara cắt cành theo công nghệ mới: Mô hình hiệu quả

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【#4】Cây Lan Vanilla Atropogon Schuit., Aver. & Rybkova, Sp. Nov

Tiến sĩ Leonid Averyanov vừa gửi cho chúng tôi tài liệu của Orchideen Journal Volume 1-1 2013 có bài của Prague Botanic Garden do Schuiteman, Avereryanov và Rybkova nói về cây Vanilla atropogon mọc tại Hòn Bà, Khánh Hòa. Tên cây lan này được đặt theo ý nghĩa của của tiếng La Tinh Atro có nghĩa là mầu đậm và podon có nghĩa là râu ở lưỡi hoa. (From atro-, dark, and pogon, beard, referring to the dark, hairy lip)

Cây lan Vanilla atropogon Schuit., Aver. & Rybkova, sp. nov

Đây là một giống phong lan hay thạch lan thân dài tới 15 thước. Thân to ngang 5-9 ly, có những đốt dài 3.5- 9.5 phân. Rễ to ngang 1.5 ly, lá dài 12-17 phân, rộng 4.2-4.7 phân, những lá ở gần ngọn nhỏ hơn. Chùm hoa dài 10-15 phân có từ 10-23 hoa, nở dần dần to 4.5 phân có mùi hôi. Hoa nở vào tháng giêng cho tới tháng 5. Lan mọc trên cao độ 180-300 th gần giòng suối, nhưng rất hiếm.

Các khoa học gia kể trên cũng xem xét đến mẫu vật do Hồ ngọc Quỳnh, thuộc khu bảo tồn Hòn Bà thu thập vào tháng 4 năm 2012 và nhận thấy rằng cây Vanilla atropogon rất giống với cây Vanilla borneensis do Rolfe mô tả, nhưng khác về mầu sắc của lưỡi hoa và mùi hôi lôi cuốn lũ ruồi nhặng đến để giúp cho cây thu phấn.

Bài viết tuy không nói rõ, nhưng nhìn vào hình ảnh cây lan do anh Hồ ngọc Quỳnh đã tìm thấy qua bài viết của Tiến sĩ Đinh Quang Diệp nói về cây Vanilla shenzhenicaZ.J. Liu & S.C. Chen, chúng tôi nghĩ rằng cây này với cây Vanilla atropogon cũng chỉ là một.

Trong bài “Cây Vanilla mới của Việt Nam” viết vào tháng 5-1012 chúng tôi đã phân tách là cây lan do anh Hồ Ngọc Quỳnh tìm thấy không phải là cây Vanilla shenzhenica mọc ở Quảng Đông vì hoa của cây này chỉ nở cum cúp chứ không mở rộng, măc dầu nguồn Wikipedia có nói: Vanilla shenzhenica là một loài phong lan thuộc chi Vanilla, Họ Phong lan. Loài lan này được mô tả khoa học lần đầu vào năm 2007 tại Thẩm Quyến, Trung Quốc. Loài này cũng hiện diện tại Hòn Bà, Khánh Hòa, Việt Nam. Những gì do nguồn Wikipedia cung cấp khó lòng có thể tin cậy được.

Như vậy loài lan Vanilla tại VN sẽ có thêm một cây lan mới: Vanilla atropogon và sẽ nâng tổng số lan ở Việt Nam lên tới 165 loài (genus) và 1117 giống (species), bởi vì trong thời gian vừa qua có tới 7 cây lan mới tìm thấy tại Hòn Bà.

Tuy nhiên trong số này còn có nhiều cây ở trong tình trạng một cây mà có 2-3 tên vì các khoa học gia đặt tên khác nhau và chưa có cơ quan nào sắp đặt lại thứ tự: tên chính thức và tên đồng danh (synonym). Chúng tôi chỉ là người chơi lan tài tử ghi nhận theo tài liệu hình ảnh và tường thuật với sự nhận xét với kiến thức hạn hẹp của mình, nhưng không thể xác định rõ ràng được.

【#5】Dendrobium Jaintianum Sabap. 2007 Có Phải Là Phong Lan

Trên Internet có một bài viết vào ngày 8-2-2013 nói về cây Dendrobium chapaense được xác nhận là cây Dendrobium jaintianum. Nguyên văn: Dendrobium chapaense confirmed to be Dendrobium jaintianum.

Tại sao có chuyện lạ như vậy? bởi vì cây Dendrobium chapaense, một cây lan đặc hữu của Việt Nam do Tiến sĩ Nga sô Leonid Averyanov tìm thấy tại Sapa, Lào Cai vào năm 2006, theo The Plant list bao gồm các tổ chức khoa học danh tiếng trên thế giới như: Kew Royal Botanical Garden, New York Botanical Garden, Global Compositae Checklist v.v… đều đã chấp nhận tên này.

Dendrobium jaintianum Sabap. 2007 có phải là Phong lan

Nhưng đọc kỹ mới biết rằng: Ngày 1-11-2013 cây lan Dendrobium chapaense của vườn lan Marlow Orchids đã được trao giải thưởng CBR/AOS (Certificate of Botanical Recognition), nhưng sau đó nhóm chuyên gia nhận diện hoa lan của Hội Hoa Lan Hoa Kỳ (The Species Identification Task Force – SITF) nhận thấy cây lan hình dưới đây không phải là Dendrobium chapaense mà là cây Dendrobium jaintianum mọc tại Ấn Độ cũng có hoa và hình dáng tương tự.

Chúng tôi nhận thấy 3 cây này có vẻ giống nhau, nhưng có một vài điểm khác biệt, nên xin trình bầy hình ảnh và tài liệu của cả 3 cây để dễ dàng nhận xét:Ngoài ra trên Internet cũng có nhiều người nhầm lẫn và có nhiều nghi vấn về những cây Den. chapaense, Den. jaintianum và Den. longicornu.

Dendrobium longicornu Lindley 1828

Mô tả: Phong lan, thân cao 30-40 cm, phủ đầy lông đen, lá 5-11 chiếc. Hoa 1-3 chiếc to 3,5-4 cm lâu tàn và thơm, nở vào mùa Thu.

Nơi mọc: Sơn La, Lai Châu, Lào Cai, Tam Đảo, Vĩnh Phú.

Ảnh: Orchid Species Ảnh: Nguyễn Minh Đức

Dendrobium chapaense Aver. 2006

Mô tả: Phong lan thân cao từ 15-25 cm, lá 7-8 chiếc, mọc hai bên dài 6-10 cm, rộng 1-1.5 cm. Chùm hoa mọc gần đỉnh dài từ 1-2.5 cm, hoa 2-5 chiếc to 2-3.5 cm. Nở vào tháng 10 và 11 không thơm. Rất giống với Den. Longicornu.

Nơi mọc: Lào Cai, Sapa.

Ảnh: Leonid Averyanov Ảnh: Chu Xuân Cảnh

Dendrobium jaintianum Sabap. 2007
Ảnh: Piotr Markiewicz

Đem sự thắc mắc này hỏi nhiều khoa học gia và dưới đây là lời giải đáp của Tiến sĩ Ron MacHatton Giám đốc Giáo Dục và Điều Hành tờ Orchids Bulletin của Hội Hoa Lan Hoa Kỳ mà chúng tôi cho là rõ ràng hơn cả:

On Tuesday, February 4, 2014 8:10 AM,

The three species, longicornu, chapaense and jaintianum are very close morphologically. Dendrobium chapaense is extremely rare in its habitat and locally endemic to one small population in Vietnam. Dendrobium jaintianum, widespad through northern India, Nepal, Bhutan and possibly further east has long been confused with the other two so you have to be very careful looking at pictures people have Vitagrowthheight.com are the distinguishing difference:

In jaintianum, the lip is, when flattened, rhombic lip without distinct sidelobes. In chapaense, the lip is distinctly trilobed and the middle lobe is a sharply pointed small triangle – this is the single easiest way to tell them apart. In jaintianum, the lip edge is serrated across most of the apical edge of the lip while in chapaense, the side lobes are completely smooth and it’s only this little jutting triangle that is serrated or ruffled. Lastly, the petals of jaintianum are much broader than those of chapaense. That one is hard without two flowers to look at but the petals of chapaense are very narrow, narrowest at the bottom and tapered near the end to a point. Those of jaintianum are widest near the middle.

I hope this helps. We, SITF, struggled with the chapaense we had to verify and had to dig back into obscure Indian journals to get to the bottom of it. It appears that the vast majority of chapaense in cultivation (if not all) are really jaintianum or longicornu.

Rất tiếc tôi không rành về từ ngữ thảo mộc bằng tiếng Việt, e rằng dịch ra chẳng ai hiểu được nên đành để y nguyên văn.

Theo Gunnar Seidenfaden, một thảo mộc gia danh tiếng đã quá cố thì mầu sắc của bông hoa không phải là yếu tố chính để phân biệt, mà chính là cấu trúc của hoa và thân lá.

Nhiều lần lướt qua trên Internet, thấy có hình của anh Chu xuân Cảnh về 2 cây này ở đâu đó, hỏi lại, anh cho biết:

Về cơ bản thì cháu thấy có 2 loại cây tương đối là khác nhau, một loại là thân mảnh và dài Den. jaintianum (trung bình khoảng 50-60 cm) và một loại thân ngắn và mập hơn Den. chapaense (chỉ khoảng 20 cm). Cả hai loại thân đều có lông đen và rất khó trồng. Hoa thì kết cấu giống nhau chỉ khác một chút về mầu sắc của lưỡi hoa (như ảnh hoanglong đưa).Cháu đã gặp cả 2 loại trên ngoài tự nhiên trên khu vực đỉnh đèo Ô Quy Hồ ở độ cao khoảng 2100 m.

Den. chapaense

Chu xuân Cảnh

Den. jaintianum

Chu xuân Cảnh

Tổng kết các phần ở trên chúng ta có 3 cây lan: Den. chapaense, Den. jaintianum và Den. longicornu với những nhận xét như sau:

• Den. chapaense đặc hữu của VN hơi giống với Den. longicornu về thân lá và mùa hoa nở.

• Den. chapaense và Den. longicornu thân ngắn (20-30 phân) còn Den. jaintianum của Ấn, Hồi thân mảnh mai và dài 50-80 phân. Bà Mani Turkel cho biết cây Den jaintianum của bà dài tới 32” (80 cm).

Nếu kể tính thêm cây Den. jaintianum, số Dendrobium tại VN sẽ là 133 giống và tổng số lên tới 1156 giống cho toàn thể loài lan.• Luỡi hoa của Den. jaintianum lớn, tròn và mầu sắc đẹp hơn là Den. chapaense

• Nhìn vào hình dáng lưỡi hoa, thấy cây Dendrobium longicornu có chiếc lưỡi hoa nhiều tua dài có những sọc mầu vàng nhạt, khác hẳn với cây Den chapense có lữoi vàng nhạt nhưng không có tua, hay thành một đường viền đậm ở lữoi hoa như cây Den. jaintianum

【#6】Một Số Loại Lan Dendrobium Rừng Nên Trồng

Phong lan trong tự nhiên có rất nhiều loài, mỗi loài có một môi trường sống khác nhau, hôm nay tôi sẽ giới thiệu với bạn một số loài lan dendrobium phổ biến dễ trồng và có hoa đẹp mà người nuôi trồng hoa phong lan nghiệp dư nên trồng:

– Dendrobium rụng lá mùa thu:

Den. anosmum (Phi điệp, Dã hạc), Dendrobium secundum (Hoàng thảo Báo hỷ),  Den. aphyllum (hạc vĩ), Den. primulinum (Long Tu), Den. wardianum (Ngũ tinh), Dendrobium heterocarpum (Hoàng thảo Lụa Vàng), Den devonianum (Hoàng thảo Tam bảo sắc), Dendrobium crystallinum (Hoàng thảo Ngọc thạch 3 màu)

– Dendrobium xanh lá quanh năm như:

Den. amabile (thủy tiên tím), Den. farmeri (Thủy tiên vuông), Den. findlayanum (Hoàng thảo Chuỗi ngọc Điện Biên), Den. chrysanthum (Phi điệp vàng), Den. lindleyi (Tiểu điệp), Den. chrysotoxum (Kim Diệp), Dendrobium draconis (Hoàng thảo Nhất điểm hồng), Dendrobium cariniferum (Hoàng thảo Nhất điểm hoàng), Dendrobium sulcatum (Thủy tiên dẹt)

Các bạn có thể chọn một trong số các loại dendrobium nêu trên mua giống và trồng cho mình một chậu lan dendrobium, chúc các bạn thành công ./.

【#7】Cấy Mắt Ngủ – Chiết Tách Lan Hồ Điệp

Cây tạo ra các mắt “ngủ” để chắc chắn rằng cây có thể sống còn, nếu chồi ngọn chết đi (bị sâu bọ ăn,…). Một thời gian sau, khi chồi ngọn phát triển, nó sản xuất chất điều hoà sinh trưởng (hormone – kích thích tố) – ngăn cản sự phát triển của các chồi khác trên thân. Nếu chồi ngọn chết, chất điều hoà sinh trưởng thiếu hụt và các chồi “ngủ” bắt đầu phát triển.

Mắt ngủ (node) là gì?

Bạn có thể tìm mắt ngủ ở đâu?

  • trên các cành hoa Lan Hồ điệp Phalaenopsis, Doritis và Lan Hạc đính.
  • trên các giả hành của Dendrobium.
  • trên các giả hành của Cattleya…
  • Trồng mắt ngủ trong đất

Chuẩn bị các mắt ngủ: Chúng ta thử làm theo kỹ thuật sau với lan Hạc đính tankervilleae và nó có kết quả rất tốt. Một trong những người tham gia khóa học đã nhân giống thành công Lan Hồ điệp của anh ta trong than bùn?. Bạn có thể xem ảnh Hồ điệp con và quy trình của anh ta dưới đây:

Nhân giống Lan Hồ điệp trong than bùn?

Một trong những người tham gia khóa học đã nhân giống thành công Lan Hồ điệp trong các viên than bùn?

Quy trình Nhân giống Lan Hồ điệp trong than bùn?

• ngâm các viên than bùn dưới vòi nước.

• bỏ các viên than bùn được trải ra vào một hộp nhựa, lấp khoảng trống giữa các viên bằng cát silica và đổ ít nước + nước dừa và trong hộp nhựa.

• đặt hộp nhựa vào trong lò vi ba (we didn´t get any informations about time and intensity / chúng tôi không có bất kỳ thông tin nào về thời gian và cường độ).

• bỏ it dớn mềm vào trong cát (when it´s cold / khi hộp nguội).

• cắt cành hoa khoảng 3 cm trên và dưới mắt ngủ.

• lột bỏ các màng bao che (của mắt ngủ).

• đặt các (đoạn) mắt ngủ vào các viên than bùn.

• đậy hộp nhựa với nắp nhựa và đặt hộp dưới ánh sáng trồng (cây).

Thích hợp là các mắt ngủ – được bạn cắt 3 cm ở phía dưới và phía trên mắt trên cành hoa. Rất quan trọng với chúng ta là phải dùng dao sắc, vì nếu không mô sẽ bị tổn thương nhiều. Tiếp, bạn phải gỡ bỏ các màng bao một cách cẩn thận.

Đặt mắt ngủ trong đất:

Đặt mắt ngủ đã chuẩn bị nằm ngang trong đất, mắt ngủ phải ở điểm cao nhất. tạo ẩm tốt và đậy hộp với nắp nhựa giống như ảnh dưới đây.

Chăm sóc về sau:

Đặt hộp chứa những mắt ngủ nơi ấm, sáng và tránh ánh nắng trực tiếp. Kiểm tra mỗi 3-4 ngày để chúng có độ ẩm đủ. Cấy mắt ngủ trong ống nghiệm

Chuẩn bị mắt ngủ

Ưu điểm của kỹ thuật này là các cây mới được nhân giống (từ) cha mẹ chúng và giống cha mẹ chúng. Chúng ta dùng kỹ thuật này để nhân giống: Phalaenopsis, Doritis pulcherima, Phaius tankervilleae (Hạc đỉnh) và Chiloschista lunifera.

Mắt ngủ phù hợp là bạn cắt chéo với 1 cm phía dưới và trên của mắt trên cành hoa. Rất quan trọng với chúng ta là dùng dao rất sắc vì nếu không, các mô sẽ bị tổn thương nhiều.

Cành hoa Hồ điệp với vỏ bọc: Tiếp, bạn phải gỡ bỏ vỏ bao mắt ngủ một cách cẩn thận. (mắt ngủ) của cành hoa Hồ điệp không vỏ bao

Dung dịch (cấy) nào bạn nên dùng ?

Để khởi đầu (cho) sự tăng trưởng của “mắt ngủ” chúng ta phải dùng dung dịch gồm có cytocinins (phytohormon). Chúng ta dùng Sigma’s P6793 (Phytotechlab P793)

Chuẩn bị nơi làm: Bạn có thể dùng dụng cụ tương tự chúng tôi mô tả trong “Kỹ thuật Gieo hạt ”

Đồ dùng cần thiết:

  • Bếp lò
  • Nồi nấu nước đường kính miệng 30cm
  • Đèn cồn (dùng sát trùng dụng cụ)
  • Bao tay cao su
  • Nhíp, kẹp (inox)
  • Dao mổ (inox)
  • Cái móc lấy hạt lan (dụng cụ cấy chuyền)

Đồ dùng cần thiết khác:

  • Ống nghiệm chứa môi trường cấy
  • Bình cấy (lọ, ống nghiệm…)
  • Giấy vệ sinh (kitchen paper)
  • Cồn 70%
  • Thuốc tẩy (như là Clorox) dùng tẩy trùng.

    Lọ (hũ) có nắp vặn (như là hũ thức ăn trẻ em – babyfood jar)

Bỏ mắt ngủ vào lọ:

Ngâm các mắt ngủ đã cắt vào cồn ethanol 70% ít giây. Sau đó đặt các mắt ngủ vào trong 0,5% hydrogen peroxide (H2O2) (nước Oxi già) trong 30 phút.

Tiếp, đặt chúng vào H2O2 3% trong 15 phút. Sau 15 phút, để các mắt ngủ đã khử trùng (trong ống nghiệm) trên vỉ – nằm trong hơi nước sôi (nơi vô trùng).

Bây giờ, nhấc ống nghiệm lên và mở (nắp) ra trong hơi nước. Nắp phải đặt trong giấy vệ sinh (kitchen paper) tẩm cồn. Lấy nhíp của bạn và dùng ngọn lửa nến để khử trùng.

Chuyển các nhíp vào nơi vô trùng (hơi nước) và gắp một mắt ngủ – đang được ngâm trong dung dịch hydrogen peroxide và đặt nó với phần cuối ở phía dưới mắt ngủ vào trong dung dịch (cấy).

Kế, nhúng nhíp vào trong nước đang sôi để làm sạch tất cả dung dịch còn sót và bỏ nhíp vào trong lọ có cồn 70%. Đậy ống nghiệm lại (trong hơi nước sôi) và đặt nó đâu đó trên bàn để dán nhãn. Với ống nghiệm kế, bạn có thể tiến hành tương tự.

Gợi ý: Để tạo dung dịch khử trùng hiệu quả hơn, thêm một giọt dung dịch rửa chén vào trong nước Oxi già.

Chăm sóc sau này

Nơi – bạn cấy các mắt ngủ sáng và ấm (khoảng 20 °C). Tránh nắng trực tiếp vì (nó) sẽ làm nóng bên trong ống nghiệm nếu chúng đang ở trong nắng trực tiếp.

Chồi Chiloschista lunifera đang tăng trưởng: Bởi vì kích thước và cấu trúc của các mắt ngủ, tỷ lệ nhiễm bệnh cao hơn so với dùng cách nẩy mầm không cộng sinh (asymbiotic seed germination).

Vì vậy, rất quan trọng là chúng ta phải kiểm tra mỗi ngày trong tuần đầu tiên, nếu có bất cứ các sự nhiễm bệnh nào. Nếu bạn tìm thấy nấm hay vi khuẩn, bạn có thể thử khử trùng chúng một lần nữa.

Nhiều mắt ngủ tiết ra các hợp chất phenolic vào dung dịch (nuôi cấy) – làm đen dung dịch. Dịch rỉ sẽ làm chết các mắt ngủ của bạn nếu bạn không thay chúng bằng dung dịch (nuôi cấy) mới. Nhiều mắt ngủ ngừng tiết ra các hợp chất phenolic sau 3 hay 4 lần thay (dung dịch nuôi cấy).

Ngay khi mắt ngủ có 2 hay 3 lá, bạn nên thay dung dịch (nuôi cấy) bằng dung dịch không có chất kích thích tố (như Sigma P6668) để bắt đầu sự phát triển rễ.

Tôi có thể làm gì nếu tôi muốn (có) nhiều hơn một cây?

Nếu bạn muốn sản xuất nhiều hơn một mầm bạn nên cắt trên 1/3 của mắt ngủ. Điều này làm mắt ngủ phát triển đến khoảng hàng tá chồi thay vì một (chồi).

Các chất điều hoà tăng trưởng (hormones)

Các chất điều hoà tăng trưởng là gì?

Các chất điều hoà tăng trưởng là bất kỳ các hợp chất hữu cơ hay tổng hợp – tác động (đến) sự tăng trưởng và sự nhân giống. Chúng được sinh ra trong cây (như trong các chồi đang tăng trưởng) để kiểm soát sự tăng trưởng.

Auxins

Auxins tác động sự tăng trưởng tế bào, ra rễ và sự hình thành chồi bất định. Chúng ngăn sự phát triển các chồi nhánh (là chồi lựa chọn cho việc đảm bảo sự ổn định di truyền). Auxins thường được dùng trong dung dịch nuôi cấy mô, hoặc là phối hợp với cytokinins suốt giai đoạn nhân giống hoặc không có cytokinins cho giai đoạn ra rễ.

Name / Tên Abbreviation / Viết tắt

Indole-3-Acetic Acid IAA

Indole-3-Butyric Acid IBA

Naphthalene Acitic Acid NAA

Phenylacetic Acid PAA

Dichlorphenoxyacetic Acid 2,4D

2,4,5-Trichlorphenoxyacetic Acid 2,4,5-T

Picloram

Dicamba

p-chlorophenoxyacetic Acid CPA

Cytokinins

Name / Tên Abbreviation / Viết tắt

Benzyladenin BA6

Benzylaminopurine BAP

Pentyladenin

Dimethylallyladenin

Kinetin

Zeatin Z

Zeatinriboside ZR

Isopentenyladenine iP

Isopentenyladenosine iPA

Thidiazuron TDZ

Gibberellins

Gibberellins là một nhóm các chất liệu từ thiên nhiên – tác động đến việc phát triển tế bào và sự tăng trưởng chiều dài của thân. E. Kurasawa (người Nhật) ghi chú vào năm 1926 rằng chất bài tiết của một loại nấm (Gibberella fugikuroi) tạo kết quả là sự tăng trưởng nhanh khác thường trong các hạt giống lúa. Chất đó là acid gibberellic – sau đó được người ta đã phân lập ở trạng thái kết tinh từ cả nấm và thực vật cao hơn.

Name / Tên Abbreviation / Viết tắt

Gibberellic Acid GA

Chlorcholinchlorid CCC

【#8】Fungi And Food Spoilage, 3Rd Ed

Fungi and Food Spoilage

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John I. Pitt

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Ailsa D. Hocking

Fungi and Food Spoilage

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John I. Pitt Honorary Research Fellow CSIRO Food and Nutritional Sciences North Ryde, NSW 2113 Australia

ISBN 978-0-387-92206-5 e-ISBN 978-0-387-92207-2 DOI 10.1007/978-0-387-92207-2 Springer Dordrecht Heidelberg London New York Library of Congress Control Number: 2009920217 # Springer ScienceþBusiness Media, LLC 2009 All rights reserved. This work may not be translated or copied in whole or in part without the written permission of the publisher (Springer ScienceþBusiness Media, LLC, 233 Spring Street, New York, NY 10013, USA), except for brief excerpts in connection with reviews or scholarly analysis. Use in connection with any form of information storage and retrieval, electronic adaptation, computer software, or by similar or dissimilar methodology now known or hereafter developed is forbidden. The use in this publication of trade names, trademarks, service marks, and similar terms, even if they are not identified as such, is not to be taken as an expssion of opinion as to whether or not they are subject to proprietary rights. Printed on acid-free paper Springer is part of Springer ScienceþBusiness Media (www.springer.com)

Preface to the Third Edition

In contrast to the second edition, the third edition of ”Fungi and Food Spoilage” is evolutionary rather than revolutionary. The second edition was intended to cover almost all of the species likely to be encountered in mainstream food supplies, and only a few additional species have been included in this new edition. The third edition repsents primarily an updating – of taxonomy, physiology, mycotoxin production and ecology. Changes in taxonomy reflect the impact that molecular methods have had on our understanding of classification but, it must be said, have not radically altered the overall picture. The improvements in the understanding of the physiology of food spoilage fungi have been relatively small, reflecting perhaps the lack of emphasis on physiology in modern microbiological science. Much remains to be understood about the specificity of particular fungi for particular substrates, of the influence of water activity on the growth of many of the species treated, and even on such basic parameters as cardinal temperatures for growth and the influence of pH and pservatives. Since 1997, a great deal has been learnt about the specificity of mycotoxin production and in which commodities and products-specific mycotoxins are likely to occur. Changes in our understanding of the ecology of the included species are also in most cases evolutionary. A great number of papers have been published on the ecology of foodborne fungi in the past few years, but with few exceptions the basic ecology of the included species remains. Recent changes in our understanding of foodborne fungi include the realisation that Aspergillus carbonarius is a major source of ochratoxin A in the world food supply, that A. westerdijkiae and not A. ochraceus is the other common Aspergillus species making this toxin and that these species are responsible for ochratoxin A in foods outside the cool temperate regions, where Penicillium verrucosum is the important species. In recent years a number of new species have been found to be capable of producing aflatoxin, but the fact remains that most aflatoxin in the global food supply is produced by A. flavus and A. parasiticus. The taxonomy of Fusarium species is still undergoing major revision. However, the renaming of Fusarium moniliforme as F. verticillioides is the only change of importance here. Recent publications have improved our understanding of species – mycotoxin relationships within Fusarium.

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Among the colleagues who helped us to ppare this edition, we wish to particularly thank Dr Anne-Laure Markovina, now of the University of Sydney, who assisted in literature searches and some cultural and photographic work, and Mr N.J. Charley who has continued his excellent work of curating the FRR culture collection, on which so much of the descriptive work in this book is based.

Preface to the Third Edition

Preface to the First Edition

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Preface to the Second Edition

In planning for the second edition of ”Fungi and Food Spoilage”, we decided that the book would benefit from a larger format, which would permit improved illustrations, and from some expansion of the text, in both numbers of species treated and overall scope. These aims have been realised. The Crown Quarto size has allowed us to include substantially larger, clearer illustrations. Many new photographs and photomicrographs have been added, the latter taken using a Zeiss Axioscop microscope fitted with Nomarski differential interference contrast optics. We have taken the opportunity to include more than 40 additional species descriptions, to add a new section on mycotoxin production for each species and to update and upgrade all of the text. Since the first edition, changes in the climate for stabilising fungal nomenclature have resulted in development of a list of ”Names in Current Use” for some important genera, including Aspergillus and Penicillium. Names of species used in the second edition are taken from that list, which was given special status by the International Botanical Congress, Tokyo, 1994. Names used in this edition have priority over any other names for a particular species. Publication of a list of ”Authors of Fungal Names” (P.M. Kirk and A.E. Ansell, Index of Fungi, Supplement: 1-95, 1992) has also stabilised names of authorities for all fungal species. Abbreviations of authors’ names used in this edition conform to those recommended by Kirk and Ansell. Some progress in standardisation of methods and media has also been made, primarily through the efforts of the International Commission on Food Mycology. The first edition included some 400 references. When we began revisionary work, we felt that the number of references in the area of food mycology had probably doubled or increased by perhaps 150% during the intervening years. In fact, this second edition includes over 1900 references, almost a five-fold increase over the 1985 edition! This provides a clear indication that interest in, and study of, food mycology has greatly increased in recent years. Modern referencing systems have enabled us to expand information from tropical sources, especially in Asia and Africa, but we are conscious of the fact that treatment of fungi found in foods on a worldwide basis remains rather incomplete. We gratefully acknowledge support and assistance from colleagues who have contributed to this new edition. Ms J.C. Eyles formatted and printed the camera

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ready copy, Ms C. Heenan collated, arranged and formatted the illustrations and Mr N.J. Charley looked after the culture collection, culture growth and colony photography. Without this level of support, the book would not have been completed.

Preface to the Second Edition

Contents 1

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

1

2

The Ecology of Fungal Food Spoilage . . . . . . . . . . . . . . . . . . . . . . . . . 2.1 Water Activity . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.2 Hydrogen Ion Concentration . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.3 Temperature . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.4 Gas Tension . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.5 Consistency. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.6 Nutrient Status . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.7 Specific Solute Effects . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.8 Preservatives . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.9 Conclusions: Food Preservation . . . . . . . . . . . . . . . . . . . . . . . .

3 3 4 5 7 8 8 8 9 9

3

Naming and Classifying Fungi . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.1 Taxonomy and Nomenclature: Biosystematics . . . . . . . . . . . . . 3.2 Hierarchical Naming . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.3 Zygomycotina. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.4 Ascomycotina . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.5 Basidiomycotina . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.6 The Ascomycete – Conidial Fungus Connection . . . . . . . . . . . . 3.7 Dual Nomenclature . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.8 Practical Classification of Fungi . . . . . . . . . . . . . . . . . . . . . . . .

11 11 12 12 13 15 15 15 16

4

Methods for Isolation, Enumeration and Identification . . . . . . . . . . . . 4.1 Sampling. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.2 Enumeration Techniques . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.2.1 Direct Plating. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.2.2 Dilution Plating . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.2.3 Incubation Conditions. . . . . . . . . . . . . . . . . . . . . . . . . . . 4.3 Sampling Surfaces . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.4 Air Sampling . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.5 Isolation Techniques . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.5.1 Yeasts . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.5.2 Moulds. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.5.3 Short Term Storage . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.6 Choosing a Suitable Medium . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.6.1 General Purpose Enumeration Media. . . . . . . . . . . . . . . 4.6.2 Selective Isolation Media. . . . . . . . . . . . . . . . . . . . . . . . . 4.6.3 Techniques for Yeasts . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.6.4 Techniques for Heat-Resistant Fungi . . . . . . . . . . . . . . . 4.6.5 Other Plating Techniques . . . . . . . . . . . . . . . . . . . . . . . . 4.7 Estimation of Fungal Biomass . . . . . . . . . . . . . . . . . . . . . . . . . . 4.7.1 Chitin . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.7.2 Ergosterol . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.7.3 Impedimetry and Conductimetry . . . . . . . . . . . . . . . . . .

19 19 19 20 21 22 22 23 23 23 24 24 25 26 27 30 32 33 34 34 35 37 xi

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Contents

4.7.4 Adenosine Triphosphate (ATP) . . . . . . . . . . . . . . . . . 4.7.5 Fungal Volatiles. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.7.6 Immunological Techniques . . . . . . . . . . . . . . . . . . . . . 4.7.7 Molecular Methods . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.8 Identification Media and Methods. . . . . . . . . . . . . . . . . . . . . . 4.8.1 Standard Methodology . . . . . . . . . . . . . . . . . . . . . . . . 4.8.2 Plating Regimen . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.8.3 Inoculation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.8.4 Additional Media and Methods . . . . . . . . . . . . . . . . . 4.8.5 Identification of Fusarium Species. . . . . . . . . . . . . . . . 4.8.6 Yeasts. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.9 Examination of Cultures . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.9.1 Colony Diameters . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.9.2 Colony Characters. . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.9.3 Preparation of Wet Mounts for Microscopy. . . . . . . . 4.9.4 Staining . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.9.5 Microscopes and Microscopy . . . . . . . . . . . . . . . . . . . 4.10 Preservation of Fungi . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.10.1 Lyophilisation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.10.2 Other Storage Techniques . . . . . . . . . . . . . . . . . . . . . . 4.11 Housekeeping in the Mycological Laboratory. . . . . . . . . . . . . 4.11.1 Culture Mites. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.11.2 Problem Fungi . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.11.3 Pathogens and Laboratory Safety . . . . . . . . . . . . . . . .

37 37 38 40 41 41 41 41 42 43 44 45 45 45 46 46 47 48 48 49 50 50 51 51

Primary Keys and Miscellaneous Fungi . . . . . . . . . . . . . . . . . . . . . . . . 5.1 The General Key . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.1.1 Notes on the General Key . . . . . . . . . . . . . . . . . . . . . . 5.2 Miscellaneous Fungi. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.3 Genus Acremonium Link . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.4 Genus Alternaria Nees: Fr.. . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.5 Genus Arthrinium Kunze . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.6 Genus Aureobasidium Viala and G. Boyer . . . . . . . . . . . . . . . . 5.7 Genus Bipolaris Shoemaker . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.8 Genus Botrytis P. Micheli: Fr. . . . . . . . . . . . . . . . . . . . . . . . . . 5.9 Genus Chaetomium Kunze . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.10 Genus Chrysonilia Arx . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.11 Genus Cladosporium Link . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.12 Genus Colletotrichum Corda . . . . . . . . . . . . . . . . . . . . . . . . . . 5.13 Genus Curvularia Boedijn. . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.14 Genus Drechslera S. Ito . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.15 Genus Endomyces Reess . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.16 Genus Epicoccum Link . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.17 Genus Fusarium Link . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.18 Genus Geotrichum Link: Fr.. . . . . . . . . . . . . . . . . . . . . . . . . . . 5.19 Genus Hyphopichia Arx and van der Walt. . . . . . . . . . . . . . . . 5.20 Genus Lasiodiplodia Ellis and Everh. . . . . . . . . . . . . . . . . . . . . 5.21 Genus Monascus Tiegh. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.22 Genus Moniliella Stolk and Dakin . . . . . . . . . . . . . . . . . . . . . . 5.23 Genus Nigrospora Zimm. . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

53 54 55 56 58 60 64 65 67 68 70 73 75 81 82 85 86 88 89 122 124 125 127 129 131

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5.24 5.25 5.26 5.27 5.28 5.29 5.30

Genus Pestalotiopsis Steyaert . . . . . . . . . . . . . . . . . . . . . . . . . . Genus Phoma Sacc. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Genus Stemphylium Wallr. . . . . . . . . . . . . . . . . . . . . . . . . . . . . Genus Trichoconiella B.L. Jain. . . . . . . . . . . . . . . . . . . . . . . . . Genus Trichoderma Pers. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Genus Trichothecium Link . . . . . . . . . . . . . . . . . . . . . . . . . . . . Genus Ulocladium Preuss . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

133 134 136 137 139 140 142

6

Zygomycetes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 6.1 Genus Absidia Tiegh. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 6.2 Genus Cunninghamella Matr. . . . . . . . . . . . . . . . . . . . . . . . . . . 6.3 Genus Mucor P. Micheli: Fr. . . . . . . . . . . . . . . . . . . . . . . . . . . 6.4 Genus Rhizomucor (Lucet and Costantin) Vuill. . . . . . . . . . . . 6.5 Genus Rhizopus Ehrenb.. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 6.6 Genus Syncephalastrum J. Schrot. ¨ …………………. 6.7 Genus Thamnidium Link . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

145 148 149 151 157 158 165 167

7

Penicillium and Related Genera . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 7.1 Genus Byssochlamys Westling . . . . . . . . . . . . . . . . . . . . . . . . . 7.2 Genus Eupenicillium F. Ludw. . . . . . . . . . . . . . . . . . . . . . . . . . 7.3 Genus Geosmithia Pitt . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 7.4 Genus Paecilomyces Bainier . . . . . . . . . . . . . . . . . . . . . . . . . . . 7.5 Genus Scopulariopsis Bainier . . . . . . . . . . . . . . . . . . . . . . . . . . 7.6 Genus Talaromyces C.R. Benj.. . . . . . . . . . . . . . . . . . . . . . . . . 7.7 Genus Penicillium Link. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 7.7.1 Penicillium subgenus Aspergilloides Dierckx . . . . . . . . . 7.7.2 Penicillium subgenus Furcatum Pitt. . . . . . . . . . . . . . . . 7.7.3 Penicillium subgenus Penicillium . . . . . . . . . . . . . . . . . . 7.7.4 Penicillium subgenus Biverticillium Dierckx . . . . . . . . .

169 170 175 182 183 187 188 194 196 207 223 263

8

Aspergillus and Related Teleomorphs. . . . . . . . . . . . . . . . . . . . . . . . . . 8.1 Genus Emericella Berk. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 8.2 Genus Eurotium Link: Fr. . . . . . . . . . . . . . . . . . . . . . . . . . . . . 8.3 Genus Neosartorya Malloch and Cain . . . . . . . . . . . . . . . . . . . 8.4 Genus Aspergillus Fr.: Fr. . . . . . . . . . . . . . . . . . . . . . . . . . . . .

275 279 281 292 295

9

Xerophiles . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 9.1 Genus Basipetospora G.T. Cole and W.B. Kendr. . . . . . . . . . . 9.2 Genus Chrysosporium Corda . . . . . . . . . . . . . . . . . . . . . . . . . . 9.3 Genus Eremascus Eidam . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 9.4 Genus Polypaecilum G. Sm. . . . . . . . . . . . . . . . . . . . . . . . . . . . 9.5 Genus Wallemia Johan-Olsen. . . . . . . . . . . . . . . . . . . . . . . . . . 9.6 Genus Xeromyces L.R. Fraser . . . . . . . . . . . . . . . . . . . . . . . . .

339 340 342 347 348 350 353

10

Yeasts. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

357

11

Fresh and Perishable Foods . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 11.1 Spoilage of Living, Fresh Foods . . . . . . . . . . . . . . . . . . . . . . . 11.2 Fruits . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 11.2.1 Citrus Fruits . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 11.2.2 Pome Fruits . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 11.2.3 Stone Fruits . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

383 383 383 384 385 386

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11.3

11.4 11.5 11.6

12

11.2.4 Tomatoes and other Solanaceous Fruit. . . . . . . . . . . 11.2.5 Melons and other Cucurbits . . . . . . . . . . . . . . . . . . . 11.2.6 Grapes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 11.2.7 Berries . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 11.2.8 Figs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 11.2.9 Tropical Fruit . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Vegetables . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 11.3.1 Peas . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 11.3.2 Beans . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 11.3.3 Onions and Garlic . . . . . . . . . . . . . . . . . . . . . . . . . . . 11.3.4 Potatoes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 11.3.5 Roots and Tubers . . . . . . . . . . . . . . . . . . . . . . . . . . . 11.3.6 Yams . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 11.3.7 Cassava . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 11.3.8 Leafy and other Green Vegetables . . . . . . . . . . . . . . Dairy Foods . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Meats . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Cereals, Nuts and Oilseeds . . . . . . . . . . . . . . . . . . . . . . . . . . . . 11.6.1 Wheat, Barley and Oats. . . . . . . . . . . . . . . . . . . . . . . 11.6.2 Rice . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 11.6.3 Maize . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 11.6.4 Soybeans and Mung Beans . . . . . . . . . . . . . . . . . . . . 11.6.5 Other Beans and Pulses . . . . . . . . . . . . . . . . . . . . . . . 11.6.6 Sunflower Seed . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 11.6.7 Sorghum. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 11.6.8 Peanuts. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 11.6.9 Cashews and Brazil Nuts . . . . . . . . . . . . . . . . . . . . . . 11.6.10 Almonds, Hazelnuts, Walnuts and Pecans . . . . . . . . 11.6.11 Pistachios . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 11.6.12 Copra . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

Spoilage of Stored, Processed and Preserved Foods . . . . . . . . . . . . . . . 12.1 Low Water Activity Foods: Dried Foods . . . . . . . . . . . . . . . . 12.1.1 Cereals . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 12.1.2 Flour . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 12.1.3 Pasta. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 12.1.4 Bakery Products. . . . . . . . . . . . . . . . . . . . . . . . . . . . . 12.1.5 Maize . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 12.1.6 Soybeans, Mung Beans, other Beans and Chickpeas . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 12.1.7 Nuts . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 12.1.8 Peanuts. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 12.1.9 Hazelnuts, Walnuts, Pecans and Almonds . . . . . . . . 12.1.10 Pistachio Nuts . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 12.1.11 Other Nuts . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 12.1.12 Coconut . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 12.1.13 Spices . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 12.1.14 Coffee Beans . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 12.1.15 Cocoa . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 12.1.16 Dried Meat . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

387 387 388 389 389 390 391 391 391 391 392 392 392 392 393 393 394 395 395 396 396 397 398 398 398 398 399 399 399 400 401 401 402 403 403 403 404 405 406 406 407 408 409 409 410 410 411 412

Contents

xv

12.2

Low Water Activity Foods: Concentrated Foods . . . . . . . . . . 12.2.1 Jams . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 12.2.2 Dried Fruit. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 12.2.3 Fruit Cakes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 12.2.4 Confectionery . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 12.2.5 Fruit Concentrates. . . . . . . . . . . . . . . . . . . . . . . . . . . . 12.2.6 Honey and Syrups . . . . . . . . . . . . . . . . . . . . . . . . . . . . Low Water Activity Foods: Salt Foods . . . . . . . . . . . . . . . . . . Intermediate Moisture Foods: Processed Meats . . . . . . . . . . . Heat Processed Acid Foods . . . . . . . . . . . . . . . . . . . . . . . . . . . Preserved Foods . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Cheese . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

412 412 413 414 415 415 416 416 417 418 418 419

Media Appendix . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

423

Glossary. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

427

References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

431

Index . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

503

12.3 12.4 12.5 12.6 12.7

Chapter 1

Introduction

From the time when primitive man began to cultivate crops and store food, spoilage fungi have demanded their tithe. Fuzzes, powders and slimes of white or black, green, orange, red and brown have silently invaded – acidifying, fermenting, discolouring and disintegrating, rendering nutritious commodities unpalatable or unsafe. Until recently, fungi have generally been regarded as causing only unaesthetic spoilage of food, despite the fact that Claviceps purpurea was linked to human disease more than 200 years ago, and the acute toxicity of macrofungi has long been known. Japanese scientists recognised the toxic nature of yellow rice 100 years ago, but 70 years elapsed before its fungal cause was confirmed. Alimentary toxic aleukia killed many thousands of people in the USSR in 1944-1947; although fungal toxicity was suspected by 1950, the causal agent, T-2 toxin, was not clearly recognised for another 25 years. Forgacs and Carll (1952), in a prophetic article, warned of the danger from common spoilage fungi, but it was not until 1960, when the famous “Turkey X” disease killed 100,000 turkey poults in Great Britain, and various other disasters followed in rapid succession, that the Western world became aware that common spoilage moulds could produce significant toxins. Since 1960 a seemingly endless stream of toxigenic fungi and potentially toxic compounds has been discovered. On these grounds alone, the statement ”It’s only a mould” is no longer acceptable to food microbiologist, health inspector or consumer. The demand for accurate identification and characterisation of food spoilage fungi has become urgent. In the flurry of research into mycotoxins, however, it must not be forgotten that food spoilage as such

J.I. Pitt, A.D. Hocking, Fungi and Food Spoilage, DOI 10.1007/978-0-387-92207-2_1, Ó Springer ScienceþBusiness Media, LLC 2009

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the process applied. As examples we can take Polypaecilum pisce on salt fish, Xeromyces bisporus on fruit cake, Cladosporium herbarum on refrigerated meat, Zygosaccharomyces bailii in pserved juices, Z. rouxii in jams and fruit concentrates, Aspergillus flavus on peanuts, Eurotium chevalieri on hazel nuts, Penicillium roqueforti on cheeses, Byssochlamys fulva in acid canned foods . . . the list of quite specific food – fungus associations is extensive. The study of such associations is one of the more important branches of the young discipline, food mycology. This book sets out to document current knowledge on the interaction of foods and fungi, in the context of spoilage and toxicity, not food production or biotechnology. Four aspects are examined. First, ecology: what factors in foods select for particular kinds of fungi? A chapter is devoted to the physical and chemical parameters which influence the growth of fungi in foods. Second, methodology: how do we isolated fungi from foods? What are the best media to use? How do we go about identifying food spoilage fungi? Third, the commodity: what fungi are usually associated with a particular food? Here ecological factors interact to produce a more or less specific habitat. Major classes of foods and their associated spoilage fungi are described. Finally, the fungus: what fungus is that? In a series of chapters, the main food spoilage moulds and yeasts are described and keyed, together with others commonly associated with food but not noted for spoilage. Where possible, further information is

1

Introduction

given on known habitats and sources, physiology, heat resistance, etc., together with a selective bibliography. Accurate information on mycotoxin production is also included. As far as possible, the pcise terminology for fungal structures used by the pure mycologist and indeed most necessary for him has been avoided in these chapters. Some concepts and terms are of course essential: these have been introduced as needed and are listed in a glossary. The taxonomic sections of this book are designed to facilitate identification of food spoilage and common food contaminant fungi. A standardised plating regimen is used, originally developed for the identification of Penicillium species (Pitt, 1979b) and extended here to other genera relevant to the food industry. Under this regimen, cultures are incubated for 1 week at 5, 25 and 378C on a single standard medium and at 258C on two others. In conjunction with the appropriate keys, this system will enable identification of most foodborne fungi to species level in just 7 days. For a few kinds of fungi, notably yeasts and xerophiles, subsequent growth under other more specialised conditions will be necessary. Finally, this book is dedicated to the general food microbiologist. May it help to restore equilibrium and assist in continued employment, when the quality assurance manager demands: ”What is it?” . . . ”How did it get in?” . . . ”What does it do?” . . . ”How do we get rid of it?” . . . and, worst of all . . . ”Is it toxic?”

Chapter 2

The Ecology of Fungal Food Spoilage

Food is not commonly regarded as an ecosystem, perhaps on the basis that it is not a ”natural” system. Nevertheless an ecosystem it is and an important one, because food plants and the fungi that colonise their fruiting parts (seeds and fruit) have been co-evolving for millennia. The seed and nut caches of rodents have provided a niche for the development of storage fungi. Fallen fruit, as they go through the cycle of decay and desiccation, have provided substrate for a range of fungi. Humans have aided and abetted the development of food spoilage fungi through their vast and varied food stores. It can be argued, indeed, some rapidly evolving organisms, such as haploid asexual fungi, are moving into niches created by man’s exploitation of certain plants as food. Food by its very nature is expected to be nutritious: therefore, food is a rich habitat for microorganisms, in contrast with the great natural systems, soil, water and plants. Given the right physico-chemical conditions, only the most fastidious microorganisms are incapable of growth in foods, so that factors other than nutrients usually select for particular types of microbial populations. Perhaps the most important of these factors relates to the biological state of the food. Living foods, particularly fresh fruits, vegetables, and also grains and nuts before harvest, possess powerful defence mechanisms against microbial invasion. The study of the spoilage of such fresh foods is more properly a branch of plant pathology than food microbiology. The overriding factor determining spoilage of a fresh, living food is the ability of specific microorganisms to overcome defence mechanisms. Generally speaking, then, spoilage of

fresh foods is limited to particular species. Such specific relationships between fresh food and fungus are discussed in Chapter 11 and under particular species. Other kinds of foods are moribund, dormant or nonliving, and the factors which govern spoilage are physical and chemical. There are eight principal factors: (l) water activity; (2) hydrogen ion concentration; (3) temperature – of both processing and storage; (4) gas tension, specifically of oxygen and carbon dioxide; (5) consistency; (6) nutrient status; (7) specific solute effects; and (8) pservatives. Each will be discussed in turn below.

2.1 Water Activity Water availability in foods is most readily measured as water activity. Water activity (aw), is a physicochemical concept, introduced to microbiologists by Scott (1957), who showed that aw effectively quantified the relationship between moisture in foods and the ability of microorganisms to grow on them. Water activity is defined as a ratio: aw ¼ p=po ; where p is the partial pssure of water vapour in the test material and po is the saturation vapour pssure of pure water under the same conditions.

J.I. Pitt, A.D. Hocking, Fungi and Food Spoilage, DOI 10.1007/978-0-387-92207-2_2, Ó Springer ScienceþBusiness Media, LLC 2009

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Water activity is numerically equal to equilibrium relative humidity (ERH) expssed as a decimal. If a sample of food is held at constant temperature in a sealed enclosure until the water in the sample equilibrates with the water vapour in the enclosed air space (Fig. 2.1a), then aw ðfoodÞ ¼ ERH ðairÞ=100: Conversely, if the ERH of the air is controlled in a suitable way, as by a saturated salt solution, at equilibrium the aw of the food will be numerically equal to the generated ERH (Fig. 2.1b). In this way, aw can be experimentally controlled, and the relation of aw to moisture (the sorption isotherm) can be studied. For further information on water activity, its measurement and significance in foods see Duckworth (1975); Pitt (1975); Troller and Christian (1978); Rockland and Beuchat (1987). In many practical situations, aw is the dominant environmental factor governing food stability or spoilage. A knowledge of fungal water relations will then enable pdiction both of the shelf life of foods and of potential spoilage fungi. Although the water relations of many fungi will be considered inpidually in later chapters, it is pertinent here to provide an overview. Like all other organisms, fungi are profoundly affected by the availability of water. On the aw scale, life as we know it exists over the range 0.9999þ to 0.60 (Table 2.1). Growth of animals is virtually confined to 1.0-0.99 aw; the permanent wilt point of mesophytic plants is near 0.98 aw; and most microorganisms cannot

Fig. 2.1 The concept of water activity (aw) (a) the relationship between aw and equilibrium relative humidity (ERH); (b) one method of controlling aw by means of a saturated salt solution, which generates a specific ERH at a specific constant temperature

The Ecology of Fungal Food Spoilage

grow below 0.95 aw. A few halophilic algae and bacteria can grow in saturated sodium chloride (0.75 aw), but are confined to salty environments. Ascomycetous fungi and conidial fungi of ascomycetous origin comprise most of the organisms capable of growth below 0.9 aw. Fungi capable of growth at low aw, in the psence of extraordinarily high solute concentrations both inside and out, must be ranked as among the most highly evolved organisms on earth. Even among the fungi, this evolutionary path must have been of the utmost complexity: the ability to grow at low aw is confined to only a handful of genera (Pitt, 1975). The degree of tolerance to low aw is most simply expssed in terms of the minimum aw at which germination and growth can occur. Fungi able to grow at low aw are termed xerophiles: one widely used definition is that a xerophile is a fungus able to grow below 0.85 aw under at least one set of environmental conditions (Pitt, 1975). Xerophilic fungi will be discussed in detail in Chapter 9. Information about the water relations of many fungi remains fragmentary, but where it is known it has been included in later chapters.

2.2 Hydrogen Ion Concentration At high water activities, fungi compete with bacteria as food spoilers. Here pH plays the decisive role. Bacteria flourish near neutral pH and fungi cannot compete unless some other factor, such as low water

2.3 Temperature

5

Table 2.1 Water activity and microbial water relations in perspectivea aw

Perspective

Foods

1.00

Blood, plant wilt point, seawater

0.95

Most bacteria

Vegetables meat, milk fruit Bread

0.90 0.85 0.80

Staphylococcus aureus

Ham Dry salami

0.75

Salt lake Halophiles

0.70

Jams Salt fish Fruit cake Confectionery Dried fruit Dry grains

Moulds

Yeasts

Basidiomycetes Most soil fungi Mucorales Fusarium Rhizopus, Cladosporium Aspergillus flavus Xerophilic Penicillia Xerophilic Aspergilli Wallemia Eurotium Chrysosporium Eurotium halophilicum

Basidiomycetes Most ascomycetes Zygosaccharomyces rouxii (salt) Zygosaccharomyces bailii Debaryomyces hansenii

0.65 Xeromyces bisporus Zygosaccharomyces rouxii (sugar) 0.60 DNA disordered a Modified from data of J.I. Pitt as reported by Brown (1974). Water activities shown for microorganisms approximate minima for growth reported in the literature.

activity or a pservative, renders the environment hostile to the bacteria. As pH is reduced below about 5, growth of bacteria becomes progressively less likely. Lactic acid bacteria are exceptional, as they remain competitive with fungi in some foods down to about pH 3.5. Most fungi are little affected by pH over a broad range, commonly 3-8 (Wheeler et al., 1991). Some conidial fungi are capable of growth down to pH 2, and yeasts down to pH 1.5. However, as pH moves away from the optimum, usually about pH 5, the effect of other growth limiting factors may become apparent when superimposed on pH. Figure 2.2 is an impssion of the combined influence of pH and aw on microbial growth: few accurate data points exist and the diagram is schematic. For heat-processed foods, pH 4.5 is of course critical: heat processing to destroy the spores of Clostridium botulinum also destroys all fungal spores. In acid packs, below pH 4.5, less severe processes may permit survival of heat-resistant fungal spores (Section 2.3).

2.3 Temperature The influence of temperature in food pservation and spoilage has two separate facets: temperatures during processing and those existing during storage.

As noted above, heat-resistant fungal spores may survive pasteurising processes given to acid foods. Apart from a few important species, little information exists on the heat resistance of fungi. Much of the information that does exist must be interpted with care, as heating menstrua and conditions can vary markedly, and these may profoundly affect heat resistance. High levels of sugars are generally protective (Beuchat and Toledo, 1977). Low pH and pservatives increase the effect of heat (Beuchat, 1981a, b; Rajashekhara et al., 2000) and also hinder resuscitation of damaged cells (Beuchat and Jones, 1978). Ascospores of filamentous fungi are more heat resistant than conidia (Pitt and Christian, 1970; Table 2.2). Although not strictly comparable, data of Put et al. (1976) indicate that the heat resistance of yeast ascospores and vegetative cells is of the same order as that of fungal conidia. Among the ascomycetous fungi, Byssochlamys species are notorious for spoiling heat processed fruit products (Olliver and Rendle, 1934; Richardson, 1965). The heat resistance of B. fulva ascospores varies markedly with isolate and heating conditions (Beuchat and Rice, 1979): a D value between 1 and 12 min at 908C (Bayne and Michener, 1979) and a z value of 6-7C8 (King et al., 1969) are practical working ps. The heat resistance of

6

2

The Ecology of Fungal Food Spoilage

Fig. 2.2 A schematic diagram showing the combined influence of water activity and pH on microbial growth

B. nivea ascospores is marginally lower (Beuchat and Rice, 1979; Kotzekidou, 1997a). Ascospores of Neosartorya fischeri have a similar heat resistance to those of Byssochlamys fulva, but have been reported less frequently as a cause of food spoilage. Heat resistant fungi are discussed further in Chapter 4. Food products may be stored at ambient temperatures, in which case pvention of spoilage relies on other parameters, or under refrigeration, where temperature is expected to play a pservative

role. Food frozen to -108C or below appears to be microbiologically stable, despite some reports of fungal growth at lower temperatures. The lowest temperatures for fungal growth are in the range -7 to 08C, for species of Fusarium, Cladosporium, Penicillium and Thamnidium (Pitt and Hocking, 1997). Nonsterile food stored at ca. 58C in domestic refrigerators, where conditions of high humidity pvail, will eventually be spoiled by fungi of these genera. At high aw and neutral pH, psychrophilic bacteria may also be important (mostly Pseudomonas species).

Table 2.2 Comparative heat resistance of ascospores and conidiaa Survivors (%) Fungus

Spore type

Initial viable count/ml

Ascospores 5.0 102 Conidia 7.3 102 Eurotium chevalieri Ascospores 1.0 103 Conidia 8.9 102 Xeromyces bisporus Ascospores 1.0 103 Aspergillus candidus Conidia 3.8 102 Wallemia sebi Conidia 7.1 102 a Heated at temperatures shown for 10 min. Data from Pitt and Christian (1970). Eurotium amstelodami

508C

608C

708C

93 107 103 128 93 102 42

85 0.3 62 0.1 30 0 0

3 0 21 0 0.3 0 0

2.4 Gas Tension

Thermophilic fungi, i.e. those which grow only at high temperatures, are rarely of significance in food spoilage. If overheating of commodities occurs, however, in situations such as damp grain, thermophiles can be a very serious problem. Thermotolerant fungi, i.e. species able to grow at both moderate and high temperatures, are of much greater significance. Aspergillus flavus and A. niger, able to grow between ca. 8 and 458C, are among the most destructive moulds known.

2.4 Gas Tension Food spoilage moulds, like almost all other filamentous fungi, have an absolute requirement for oxygen. However, many species appear to be efficient oxygen scavengers, so that the total amount of oxygen available, rather than the oxygen tension, determines growth. The concentration of oxygen dissolved in the substrate has a much greater influence on fungal growth than atmospheric oxygen tension (Miller and Golding, 1949). For example, Penicillium expansum grows virtually normally in 2.1% oxygen over its entire temperature range (Golding, 1945), and many other common food spoilage fungi are inhibited only slightly when grown in nitrogen atmospheres containing approximately 1.0% oxygen (Hocking, 1990). Paecilomyces variotii produced normal colonies at 258C under 650 mm of vacuum (Pitt, unpublished). Most food spoilage moulds appear to be sensitive to high levels of carbon dioxide, although there are notable exceptions. When maintained in an atmosphere of 80% carbon dioxide and 4.2% oxygen, Penicillium roqueforti still grew at 30% of the rate in air (Golding, 1945), provided that the temperature was above 208C. In 40% CO2 and 1% O2, P. roqueforti grew at almost 90% of the rate in air (Taniwaki et al., 2001a). Xeromyces bisporus has been reported to grow in similar levels of carbon dioxide (Dallyn and Everton, 1969). Byssochlamys species appear to be particularly tolerant of conditions of reduced oxygen and/or elevated carbon dioxide. Growth of Byssochlamys nivea was little affected by replacement of nitrogen in air by carbon dioxide, and growth in carbon dioxide-air mixtures was proportional only to

7

oxygen concentration, at least up to 90% carbon dioxide (Yates et al., 1967). Both Byssochlamys nivea and B. fulva were capable of growth in atmospheres containing 20, 40 or 60% carbon dioxide with less than 0.5% oxygen, but inhibition increased with increasing carbon dioxide concentration (Taniwaki et al., 2001a). Byssochlamys fulva is capable of growth in 0.27% oxygen, but not in its total absence (King et al., 1969). It is also capable of fermentation in fruit products, but psumably only if some oxygen is psent. At least some species of Mucor, Rhizopus and Fusarium are able to grow and ferment in bottled liquid products and sometimes cause fermentative spoilage. Growth under these conditions may be yeast-like. Species of Mucor, Rhizopus and Amylomyces used as starter cultures in Asian fermented foods can grow under anaerobic conditions, demonstrated by growth in an anaerobe jar with a hydrogen and carbon dioxide generator (Hesseltine et al., 1985). Other authors have reported growth under anaerobic conditions of such fungi as Mucor species, Absidia spinosa, Geotrichum candidum, Fusarium oxysporum and F. solani (Stotzky and Goos, 1965; Curtis, 1969; Taniwaki, 1995). The yeast-like fungus Moniliella acetoabutans can cause fermentative spoilage under totally anaerobic conditions (Stolk and Dakin, 1966). As a generalisation, however, it is still correct to state that most food spoilage problems due to filamentous fungi occur under aerobic conditions, or at least where oxygen tension is appciable, due to leakage or diffusion through packaging. In contrast, Saccharomyces species, Zygosaccharomyces species and other fermentative yeasts are capable of growth in the complete absence of oxygen. Indeed, S. cerevisiae and Z. bailii can continue fermentation under several atmospheres pssure of carbon dioxide. This property of S. cerevisiae has been harnessed by mankind for his own purposes, in the manufacture of bread and many kinds of fermented beverages. Z. bailii, on the other hand, is notorious for its ability to continue fermenting at reduced water activities in the psence of high levels of pservatives. Fermentation of juices and fruit concentrates may continue until carbon dioxide pssure causes container distortion or explosion. The closely related species Zygosaccharomyces rouxii is a xerophile and causes

8

spoilage of low-moisture liquid or packaged products such as fruit concentrates, jams and dried fruit. The difference in oxygen requirements between moulds and fermentative yeasts is one of the main factors determining the kind of spoilage a particular commodity will undergo.

2.6 Nutrient Status As noted in the pamble to this chapter, the nutrient status of most foods is adequate for the growth of any spoilage microorganism. Generally speaking, however, it appears that fungal metabolism is best suited to substrates high in carbohydrates, whereas bacteria are more likely to spoil proteinaceous foods. Lactobacilli are an exception. Most common mould species appear to be able to assimilate any food-derived carbon source with the exception of hydrocarbons and highly condensed polymers such as cellulose and lignin. Most moulds are equally indifferent to nitrogen source, using nitrate, ammonium ions or organic nitrogen sources with equal ease. Some species achieve only limited growth if amino acids or proteins must provide both carbon and nitrogen. A few isolates classified in Penicillium subgen. Biverticillium are unable to utilise nitrate (Pitt, 1979b).

2

The Ecology of Fungal Food Spoilage

Some xerophilic fungi are known to be more demanding. Ormerod (1967) showed that growth of Wallemia sebi was strongly stimulated by proline. Xerophilic Chrysosporium species and Xeromyces bisporus also require complex nutrients, but the factors involved have not been defined (Pitt, 1975). Yeasts are often fastidious. Many are unable to assimilate nitrate or complex carbohydrates; a few, Zygosaccharomyces bailii being an example, cannot grow with sucrose as a sole source of carbon. Some require vitamins. These factors limit to some extent the kinds of foods susceptible to spoilage by yeasts. A further point on nutrients in foods is worth making here. Certain foods (or nonfoods) lack nutrients essential for the growth of spoilage fungi. Addition of nutrient, for whatever reason, can turn a safe product into a costly failure. Two cases from our own experience illustrate this point, both involving spoilage by the pservativeresistant yeast Zygosaccharomyces bailii. In the first, a highly acceptable (and nutritious) carbonated beverage containing 25% fruit juice was eventually forced from the Australian market because it was impractical to ppare it free of occasional Z. bailii cells. Effective levels of pservative could not be added legally and pasteurisation damaged its flavour. Substitution of the fruit juice with artificial flavour and colour removed the nitrogen source for the yeast. A spoilage free product resulted, at the cost of any nutritional value and a great reduction in consumer acceptance. The other case concerned a popular water-ice confection, designed for home freezing. This confection contained sucrose as a sweetener and a pservative effective against yeasts utilising sucrose. One production season the manufacturer decided, for consumer appeal, to add glucose to the formulation. The glucose provided a carbon source for Zygosaccharomyces bailii, and as a result several months production, valued at hundreds of thousands of dollars, was lost due to fermentative spoilage.

2.7 Specific Solute Effects As stated earlier, microbial growth under conditions of reduced water availability is most satisfactorily described in terms of aw. However,

2.9 Conclusions: Food Preservation

the particular solutes psent in foods can exert additional effects on the growth of fungi. Scott (1957) reported that Eurotium (Aspergillus) amstelodami grew 50% faster at its optimal aw (0.96) when aw was controlled by glucose rather than magnesium chloride, sodium chloride or glycerol. Pitt and Hocking (1977) showed a similar effect for Eurotium chevalieri and reported that the extreme xerophiles Chrysosporium fastidium and Xeromyces bisporus grew poorly if at all in media containing sodium chloride as the major solute. In contrast Pitt and Hocking (1977) and Hocking and Pitt (1979) showed that germination and growth of several species of Aspergillus and Penicillium was little affected when medium aw was controlled with glucose-fructose, glycerol or sodium chloride. Zygosaccharomyces rouxii, the second most xerophilic organism known, has been reported to grow down to 0.62 aw in fructose (von Schelhorn, 1950). Its minimum aw for growth in sodium chloride is reportedly much higher, 0.85 aw (Onishi, 1963). Some fungi are halophilic, being well adapted to salty environments such as salted fish. Basipetospora halophila and Polypaecilum pisce grow more rapidly in media containing NaCl as controlling solute (Andrews and Pitt, 1987; Wheeler et al., 1988c). Such fungi have been called halophilic xerophiles to distinguish them from obligately halophilic bacteria.

2.8 Preservatives Obviously, pservatives for use in foods must be safe for human consumption. Under this constraint, food technologists in most countries are limited to the use of weak acid pservatives: benzoic, sorbic, nitrous, sulphurous, acetic and propionic acids – or, less commonly, their esters. In the concentrations permitted by most food laws, these acids are useful only at pH levels up to their pKa plus one pH unit, because to be effective they must be psent as the undissociated acid. For studies of the mechanism of action of weak acid pservatives see Warth (1977, 1991); Brul and Coote (1999); Stratford and Anslow (1998) and Stratford and Lambert (1999).

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The use of chemical pservatives in foods is limited by law in most countries to relatively low levels and to specific foods. A few fungal species possess mechanisms of resistance to weak acid pservatives, the most notable being Zygosaccharomyces bailii. This yeast is capable of growth and fermentation in fruit-based cordials of pH 2.9-3, of 458C Brix and containing 800 mg/L of benzoic acid (Pitt and Hocking, 1997). The yeast-like fungus Moniliella acetoabutans can grow in the psence of 4% acetic acid and survive in 10% (Pitt and Hocking, 1997). Of the filamentous fungi, Penicillium roqueforti appears to be especially resistant to weak acid pservatives and this property has been suggested as a useful aid to isolation and identification (Engel and Teuber, 1978).

2.9 Conclusions: Food Preservation It is evident from the above discussion that the growth of fungi in a particular food is governed largely by a series of physical and chemical parameters, and definition of these can assist greatly in assessing the food’s stability. The situation in practice is made more complex by the fact that such factors frequently do not act independently, but synergistically. If two or more of the factors outlined above act simultaneously, the food may be safer than expected. This has been described by Leistner and Rodel (1976) as the ”hurdle concept”. ¨ This concept has been evaluated carefully for some commodities such as fermented sausages and is now widely exploited in the production of shelf stable bakery goods and acid sauces. For most fungi, knowledge remains meagre about the influence of the eight parameters discussed here on germination and growth. However, sufficient information is now available that some rationale for spoilage of specific commodities by certain fungi can be attempted, especially where one or two parameters are of overriding importance. This topic is considered in later chapters devoted to particular commodities.

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Chapter 3

Naming and Classifying Fungi

As with other living organisms, the name applied to any fungus is a binomial, a capitalised genus name followed by a lower case species name, both written in italics or underlined. The classification of organisms in genera and species was a concept introduced by Linneaus in 1753 and it is the keystone of biological science. It is as fundamental to the biologist as Arabic decimal numeration is to the mathematician. Here the analogy ends: the concept of ”base 10” is rigorous; the concept of a species, fundamental as it is, is subjective and dependent on the knowledge and concepts of the biologist who described it.

3.1 Taxonomy and Nomenclature: Biosystematics Once biologists began to describe species and to assemble them into genera, questions about their relationships began to arise: is species x described by Jones in 1883 the same as species y described by Smith in 1942? Does species z, clearly distinct from x and y in some characters, belong to the same genus? The study of these relationships is termed taxonomy. Modern taxonomy is based on sound scientific principles, but still involves subjective judgment. When the decision is made that species x and species y are the same, however, the taxonomist must follow clearly established procedures in deciding which name must be used (”has priority”). The application of these procedures is termed nomenclature and, for fungi, plants and algae, is governed by the International Code of Botanical Nomenclature (ICBN). The ICBN is a relatively complex document of about 70 Articles dealing with all aspects of

correctly naming plants, algae and fungi. It is amended every 6 years by special sessions at each International Botanical Congress and is republished thereafter. The 17th version of the ICBN (the Vienna code) is the most recently published (McNeill et al., 2006). The ICBN impinges only indirectly on the work of the practicing mycologist or microbiologist. It is nevertheless of vital importance to the orderly naming of all plant life; to ignore the ICBN is to invite chaos. Where confusion arises over the correct name for a botanical species – a constant source of irritation to the nontaxonomist – it stems usually from one of three causes: indecision by, or disagreement among, taxonomists on what constitutes a particular species; incorrect application of the provisions of the ICBN; or ignorance of earlier literature. To return to our example, when species x and species y are seen to be the same, x has priority because it was published earlier; y becomes a synonym of x. Important synonyms are often listed after a name to aid the user of a taxonomy, and this procedure has been followed here. Through ignorance, the same species name may be used more than once, for example, Penicillium thomii Maire 1915 and P. thomii K.M. Zalessky 1927. The name P. thomii has been given to two quite different fungi. Clearly P. thomii Maire has priority; the later name is not valid. To avoid ambiguity, correct practice in scientific publication is to cite the author of a species at first mention, and before any formal description. The ICBN provides rules to govern change of genus name also. In our example, if species z is transferred to the genus to which species x and y belong, it retains its species name but takes the new

J.I. Pitt, A.D. Hocking, Fungi and Food Spoilage, DOI 10.1007/978-0-387-92207-2_3, Ó Springer ScienceþBusiness Media, LLC 2009

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Naming and Classifying Fungi

genus name. The original author of the name z is placed in brackets after the species name, followed by the name of the author who transferred it to the correct genus. For example, Citromyces glaber Wehmer 1893 became Penicillium glabrum (Wehmer) Westling 1893 on transfer to Penicillium by Westling in 1911. Note the use of Latinised names: glaber (masculine) became glabrum (neuter) to agree with the gender of the genus to which it was transferred. Further points on the use of the ICBN arise from this example. P. glabrum retains its date of original publication, and therefore takes priority over P. frequentans Westling 1911 if the two species are combined. When Raper and Thom (1949) combined the two species, a taxonomically correct decision, they retained the name P. frequentans, which was nomenclaturally incorrect, causing confusion when Subramanian (1971) and Pitt (1979b) took up the correct name. It is worth pointing out that the confusion in this and similar situations arose from Raper and Thom’s action in ignoring the provisions of the ICBN, not from that of later taxonomists who correctly interpted it.

connection between the Ascomycetes, the fungi that produce sexual spores in sacks, and the Deuteromycetes, where spores are always asexual, has been known for a long time. However, molecular taxonomy has provided the fundamental assurance needed to make this change. From the point of view of the food mycologist, this is a mixed blessing. The demands of the molecular systematists may yet make the taxonomy of foodborne fungi even more complicated. The taxonomic system used here is believed to be both practical and in line with the current ”best practice” of the nomenclaturalists. The hierarchical subpisions in Kingdom Fungi of interest in the psent context are shown below, using as examples three genera and species important in food spoilage:

3.2 Hierarchical Naming

Note that names of genera, species and varieties are italicised or underlined, while higher taxonomic ranks are not. Three subkingdoms of the kingdom Fungi include genera of significance in food spoilage. As indicated in the examples above, these are Zygomycotina, Ascomycotina and (much less commonly) Basidiomycotina. Fungi from each of these subkingdoms have quite distinct properties, shared with other genera and species from the same subkingdom. Unlike other texts, this book will not rely on initial recognition of a correct subkingdom before identification of genus and species can be undertaken. Nevertheless, identification of the subkingdom can provide valuable information about a fungus, so the principal properties of these three subkingdoms are described below.

A given biological entity, or taxon in modern terminology, can be given a hierarchy of names: a cluster of related species is grouped in a genus, of related genera in families, of families in orders, orders in classes, and classes in subkingdoms. Similarly a species can be pided into smaller entities: subspecies, varieties and formae speciales (a term usually reserved for plant pathogens). In most modern classifications, the fungi are ranked, like plants and animals, as a separate kingdom. Traditionally, fungi have been pided into several subkingdoms, based on spore type and some environmental considerations. Modern molecular methods have revolutionised this. Fungi have been shown to be more closely related to animals than plants, where traditional taxonomy has always placed them. Some of the so-called ”lower fungi” have been shown not to be fungi of all (though mycologists will no doubt continue to study them). The most important change from the point of view of the food mycologist is the demise of the subkingdom Deuteromycotina, and its absorption (almost entirely) into the Ascomycotina. The

Kingdom Subkingdom Class Order Family Genus Species Variety

Fungi Zygomycotina Zygomycetes Mucorales Mucoraceae Rhizopus stolonifer

Fungi Ascomycotina Plectomycetes Eurotiales Trichocomaceae Eurotium chevalieri intermedius

Fungi Basidiomycotina Wallemiomycetes Wallemiales Wallemiaceae Wallemia sebi

3.3 Zygomycotina Most fungi within the subkingdom Zygomycotina belong to the class Zygomycetes. Fungi in this class possess three distinctive properties:

3.4 Ascomycotina

1. Rapid growth. Most isolates grow very rapidly, often filling a Petri dish of malt extract agar with loose mycelium in 2-4 days. 2. Nonseptate mycelium. Actively growing mycelia are without septa (cross walls) and are essentially unobstructed. This allows rapid movement of cell contents, termed ”protoplasmic streaming”, which can be seen readily by transmitted light under the binocular microscope. In wet mounts the absence of septa is usually obvious (Fig. 3.1a). 3. Reproduction by sporangiospores. The reproductive structure characteristic of Zygomycetes is the sporangiospore, an asexually produced spore which in genera of interest here is usually produced inside a sac, the sporangium, on the end of a long specialised hypha. Sporangiospores are produced very rapidly. From the food spoilage point of view, the outstanding properties of Zygomycetes are very rapid growth, especially in fresh foods of high water activity; inability to grow at low water activities (no Zygomycetes are xerophiles); and lack of resistance to heat and chemical treatments. From the food safety point of view, Zygomycetes have rarely been reported to produce mycotoxins.

3.4 Ascomycotina The subkingdom Ascomycotina is distinguished from Zygomycotina by a number of fundamental characters, the most conspicuous being the production of septate mycelium (Fig. 3.1b). Consequent on

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this, growth of fungi in this subkingdom is usually slower than that of Zygomycetes, although there are some exceptions. Fungi in the subkingdom Ascomycotina, loosely called ”ascomycetes”, characteristically produce their reproductive structures, ascospores, within a sac called the ascus (plural, asci, Fig. 3.2a, b). In most fungi, nuclei normally exist in the haploid state. At one point in the ascomycete life cycle, diploid nuclei are produced by nuclear fusion, which may or may not be pceded by fusion of two mycelia. These nuclei undergo meiosis within the ascus, followed by a single mitotic pision and then differentiation into eight haploid ascospores. In most genera relevant to this work, asci can be recognised in stained wet mounts by their shape, which is spherical to ellipsoidal and smoothly rounded; size, which is generally 8-15 mm in diameter; and the psence when maturity approaches of eight ascospores tightly packed within their walls. At maturity asci often rupture to release the ascospores, which are thick walled, highly refractile, and often strikingly ornamented (Fig. 3.2c, d). Two other characteristics of asci are significant: generally they mature slowly, after incubation for 10 days or more at 258C, and they are usually borne within a larger, macroscopic body, the general term for which is ascocarp. Genera of interest here usually produce asci and ascospores within a spherical, smooth-walled body, the cleistothecium (Fig. 3.3a), or a body with hyphal walls, the gymnothecium (Fig. 3.3b). Ascospores are highly condensed, refractile spores, which are often resistant to heat, pssure

Fig. 3.1 (a) Nonseptate mycelium of Syncephalastrum racemosum; (b) septate mycelium of Fusarium equiseti

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Naming and Classifying Fungi

Fig. 3.2 Asci and ascocarps: (a) asci of Talaromyces species; (b) asci of Byssochlamys fulva; (c) ascospores of Eupenicillium alutaceum; (d) ascospores of Neosartorya quadricincta. Bars ¼ 5 mm

and chemicals. Almost all xerophilic fungi are ascomycetes. Besides their sexual spores, ascospores, ascomycetes commonly produce asexual spores. Formed after mitotic nuclear pision, these spores are borne singly or in chains, in most genera of interest here from more or less specialised hyphal structures. The general term for this type of spore is conidium (plural, conidia), but other more specialised terms

exist for specific kinds of conidia. Along the evolutionary process, some Ascomycetes with welldeveloped asexual stages lost the ability to produce ascospores, and rely entirely on conidia for dispersal. Conidia, and the specialised hyphae from which they are borne, are astonishingly perse in appearance. The size, shape and ornamentation of conidia and the complexity of the structures producing them

Fig. 3.3 (a) Cleistothecia of Eupenicillium; (b) gymnothecia of Talaromyces. SEM. Bars = 50 mm

3.7 Dual Nomenclature

provide the basis for classification of Ascomycetes that no longer produce the ascosporic (sexual) stage. Lacking ascospores, conidial fungi are not usually heat resistant, but conidia may be quite resistant to chemicals. Some conidial fungi are xerophilic.

3.5 Basidiomycotina The Subkingdom Basidiomycotina includes mushrooms, puffballs and the plant pathogenic rusts and smuts. Until recently it was not considered of any interest to the food mycologist. However, molecular studies indicate that the small brown species Wallemia sebi, long a curiosity because of its lack of resemblance to any other fungus, is a basidiomycete. It has no obvious phylogenetic affinity with any other genus and has now been classified in its own order, Wallemiales (Zalar et al., 2005). Only one other species of foodborne fungi, Trichosporonoides nigrescens Hocking and Pitt (1981), has a known affinity with this subkingdom.

3.6 The Ascomycete – Conidial Fungus Connection It was established more than a century ago that many fungal species carry the genetic information to produce both ascospores and conidia. These two kinds of spores are produced by different mechanisms and have different functions, so they are not always formed simultaneously. Not surprisingly, mycologists sometimes have given different generic and species names to a single fungus producing both an ascosporic and a conidial state. The usage of these names under the ICBN depends on the circumstances under which they were originally given. Some of these circumstances are discussed briefly below. The ascomycete state, now usually referred to as the teleomorph, is regarded by nomenclaturalists as the more important reproductive state, and the name applied to the teleomorph should be used when the ascomycete state is psent. If the conidial state is also in evidence, the fungus is now a holomorph and is still correctly known by the teleomorph name. If the conidial state, known as the

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anamorph, has a separate name, this strictly speaking applies to the conidial state. It should be used only when the ascomycete state is absent, or to refer specifically to the conidial state if the ascomycete is psent. However, the reader is warned that some anamorphic names are, and will continue to be, in common use for holomorphic fungi. Under the Articles of the ICBN, a generic name originally given to an anamorphic or conidial fungus cannot be used for a teleomorphic or ascomycetous fungus. For example, the name Penicillium, originally given to an anamorphic fungus with no known teleomorph, cannot be used for the teleomorphs later found to be produced by other Penicillium species. Such teleomorphs are classified in the genera Eupenicillium or Talaromyces, depending on whether ascospores are produced in cleistothecia or gymnothecia. Correct species names for the ascomycetous and conidial states of a single holomorphic fungus may or may not be the same, depending both on the circumstance in which the names were originally given, and on later synonymy. For example, Eupenicillium ochrosalmoneum Scott and Stolk and Penicillium ochrosalmoneum Udagawa refer to the teleomorph and anamorph of a single fungus. Udagawa (1959) described the anamorph; the teleomorph was later found, in the same isolate, by Scott and Stolk (1967). On the other hand, the anamorph of Eupenicillium cinnamopurpureum Scott and Stolk (1967) is Penicillium phoeniceum van Beyma (1933), with P. cinnamopurpureum Abe ex Udagawa (1959) as a synonym. Scott and Stolk (1967) found a teleomorph in Udagawa’s P. cinnamopurpureum; Pitt (1979b) later showed that this species was a synonym of the earlier P. phoeniceum. E. cinnamopurpureum, the first name applied to the teleomorph, is unaffected by this change in the anamorph name. In passing, note that ”Abe ex Udagawa” indicates invalid (incomplete) publication of this species by Abe, with validation later by Udagawa. The species dates from the year of validation.

3.7 Dual Nomenclature An important point here is that some isolates of Penicillium phoeniceum regularly produce the teleomorphic state Eupenicillium cinnamopurpureum, while

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others, taxonomically indistinguishable, fail to produce a teleomorph at all. Because of this, it is essential to have a separate name for teleomorph and anamorph. The system of two names for a single fungus, known as dual nomenclature, has a place in the classification of fungi despite its apparent complexity. In the descriptions in later chapters, fungi for which both teleomorphs and anamorphs are known have both names listed. As noted above, if both states are found in a particular isolate, the teleomorph name is the more appropriate: to use that given to the anamorph is not incorrect, but this name is more sensibly applied to the conidial state only. Dual nomenclature would be relatively simple if the relationship between anamorph and teleomorph was always one to one. This is not the case. As has already been mentioned, species classified in Penicillium may produce teleomorphs in two genera, Eupenicillium and Talaromyces. On the other hand Talaromyces produces anamorphs in two genera, Penicillium and Paecilomyces. Aspergillus is the anamorph of eight or ten teleomorphic genera. Most teleomorph-anamorph relationships encountered in food mycology belong to the genera mentioned here. These relationships will be described where necessary under these particular genera.

3.8 Practical Classification of Fungi Fungi are classified in a vast array of orders, families, genera and species. Among natural organisms, the numbers of taxa of fungi are rivalled only by those of the flowering plants and insects. Estimates of fungal species range as high as 1.5 million; only 5% of this number have so far been described (Hawksworth, 1991). Many fungi are highly specialised. Some will grow only in particular environments such as soil or water; many are obligate parasites and require a specific host, such as a particular plant species, and will not grow in artificial culture; many grow only in association with plant roots. From the point of view of the food microbiologist, these kinds of fungi are irrelevant. In one sense, most fungi which spoil foods are also highly specialised, their speciality being the ability to obtain nutrients from, and hence grow on, dead, dormant or moribund plant material more or less regardless of source. The

3

Naming and Classifying Fungi

principal factors influencing food spoilage by fungi are physico-chemical and have already been outlined in Chapter 2. The point being made here is that food spoilage fungi are classified in just a few orders and a relative handful of genera. For this reason there is much to be said for food mycologists avoiding the use of a traditional, hierarchical classification as outlined above and employing a less formal approach to the identification of the fungi of interest to them. In the psent work, this pragmatic approach has been followed as far as possible:

The use of specialised terms has been kept to a

minimum, while being cognisant of the need for clarity of expssion. Hierarchical classification has been avoided as far as possible, consistent with retaining a logical approach to the psentation of fungi which are related or of similar appearance. Identification procedures used have been designed to be simple and comphensible, avoiding the use of specialised equipment or procedures unavailable in the routine laboratory. To this end, identification of nearly all species included in this work is based entirely on inoculation of a single series of Petri dishes, incubation under carefully standardised conditions and examination by traditional light microscopy. A standard plating regimen has been used for the initial examination of all isolates (except yeasts), so that identification procedures can be carried out without foreknowledge of genus or even subkingdom. Cultural characters, which can be broadly defined as the application of microbiological techniques to mycology, have been used throughout.

The use of cultural characters has long been implicit in the study of fungi in pure culture on artificial substrates, especially in such genera as Aspergillus and Penicillium, genera of paramount importance in food spoilage. In Penicillium, cultural characters have been used as taxonomic criteria since the turn of the 20th century, but have assumed greater importance through the work of Pitt (1973, 1979b), who used the measurement of colony diameters, following incubation under standardised conditions, as a taxonomic criterion. The use of

3.8 Practical Classification of Fungi

pure culture techniques and growth data in fungal taxonomy is now widespad. Food microbiologists, the primary audience for this book, are familiar with cultural techniques and the use of a wide range of media and varied incubation conditions, so the authors make no apology for the taxonomic approach used in the psent work. This approach is a logical extension of the system used by the first author in Penicillium taxonomy and which has been found to have a much broader applicability. In the field of mycology, different genera have been studied by many different people of varied backgrounds and for different reasons. Consequently, keys and descriptions have been based on a wide variety of media, often traditional formulations incorporating all sorts of natural products. This heterogeneity makes comparisons difficult

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and adds unnecessary complexity to the task of the nonspecialist confronted with a range of fungal genera. The approach used here has been to examine every isolate (excluding yeasts) by a single system: inoculation onto a standard set of Petri dishes and examination of them culturally and microscopically after 7 days incubation. Most of the genera and species included in this book can be identified immediately, at that point. Only in exceptional cases has it been found necessary to reinoculate isolates onto a further set of media in order to complete identification. The exceptional fungi are first the xerophiles, many of which grow poorly if at all on the standard media, and, second, genera such as Fusarium and Trichoderma, in which some species cannot readily be differentiated on the standard regimen. Details of the techniques used are given in Chapter 4.

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Chapter 4

Methods for Isolation, Enumeration and Identification

This chapter describes techniques and media suitable for the enumeration, isolation and identification of fungi from foods. Some techniques are similar to those used in food bacteriology; others have been developed to meet the particular needs of food mycology. Most of the media have been specifically formulated for foodborne fungi. The approach taken here is designed to provide a systematic basis for the study of food mycology. In 1984 a group of about 30 of the world’s foremost scientists in food mycology met in Boston, Massachusetts, USA, to hear and discuss a wide range of psentations that explored many aspects of methodology in food mycology. Agreement was reached on broad issues and areas requiring further work pinpointed. The proceedings were published as ”Methods for the Mycological Examination of Food” (King et al., 1986). At a second workshop, held in Baarn, the Netherlands, in 1990, results of a number of collaborative studies on media and methods were psented and some standardised protocols developed. The proceedings, published as ”Modern Methods in Food Mycology” (Samson et al., 1992), provided a comphensive overview of current thinking in this field. The working group which organised those two workshops was then formalised as the International Commission on Food Mycology (ICFM), a commission under the auspices of the Mycology Division of the International Union of Microbiological Societies (IUMS). ICFM is dedicated to international standardisation of methods in food mycology. Subsequent ICFM workshops were held in Copenhagen, Denmark (1994), Uppsala, Sweden (1998), Samsø, Denmark (2003) and Key West,

Florida (2007). Papers from the third and fourth workshops were published in the International Journal of Food Microbiology and the proceedings of the fifth (Samsø) workshop were published as ”Advances in Food Mycology” (Hocking et al., 2006a). The methodology described below is based on recommendations from ICFM and repsents current thinking within the food mycology community. However, no formal endorsement from ICFM is implied.

4.1 Sampling It must be emphasised at the outset that results from mycological assays of foods are only as good as the samples used. However, sampling is beyond the scope of this text. Excellent treatises on sampling plans for food bacteriological purposes have been produced by the International Commission on Microbiological Specifications for Foods (ICMSF, 1986, 2002) and are generally applicable to food mycology.

4.2 Enumeration Techniques Quantification of the growth of filamentous fungi is more difficult than for bacteria or yeasts. Vegetative growth consists of hyphae, which are not readily detached from the substrate and which survive blending poorly. When sporulation occurs,

J.I. Pitt, A.D. Hocking, Fungi and Food Spoilage, DOI 10.1007/978-0-387-92207-2_4, Springer ScienceþBusiness Media, LLC 2009

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very high numbers of spores may be produced, causing sharp rises in viable counts, often without any great increase in biomass. The estimation of fungal growth or biomass is not easy, because no primary standard exists (such as cell numbers used for yeasts and bacteria). Although techniques for quantifying biomass have improved in recent years, most food laboratories continue to rely on viable counting (dilution plating) for detecting and quantifying fungal growth in foods. As well as dilution plating, a second standard method, known as direct plating, has been developed for estimating fungal numbers and growth in foods. Both methods are described in detail below. Techniques for biomass estimation will be discussed later.

4.2.1 Direct Plating Contributions at the international workshops mentioned above emphasised the use of direct plating as the pferred method for detecting, enumerating and isolating fungi from particulate foods such as grains and nuts. In direct plating, food particles are placed directly on solidified agar media. In most situations, particles should be surface disinfected before plating, as this removes the inevitable surface contamination arising from dust and other sources and permits recovery of the fungi actually growing in the particles. This process provides an effective measure of inherent mycological quality and permits assessment of the potential psence of mycotoxins as well. Surface disinfection should be omitted only where surface contaminants become part of the downstream mycoflora, for example, in grain intended for flour manufacture. Even here, surface disinfection before direct plating provides the most realistic appraisal of actual grain quality. Results from direct plating analyses are expssed as percentage infection of particles. The technique provides no direct indication of the extent of fungal invasion in inpidual particles. However, it is reasonable to assume that a high percentage infection is correlated with extensive invasion in the particles and a higher risk of mycotoxin occurrence.

4 Methods for Isolation, Enumeration and Identification

The standard protocol recommended by the ICFM (Hocking et al., 2006a, p. 344) is given below, with amplification where necessary. Surface disinfection. Surface disinfection is carried out by immersing particles in a chlorine solution. Household chlorine bleach, nominally 4-5% active chlorine, is effective. Dilute the chlorine 1 to 10 with water before use, to provide an approximately 0.4% solution. Immerse particles for 2 min, stirring occasionally, then drain the chlorine. Chlorine solutions are rapidly denatured by organic matter, so it is important to use a surplus of chlorine solution (10 times the volume of the particles) and to use the solution only once. This process is conveniently carried out in 250-500 ml beakers. Place 50 or more particles in the beaker and add chlorine. To dislodge air bubbles, immediately stir with a pair of forceps, leaving them in the solution, and pferably cover the beaker with a watch glass. The watch glass simplifies decanting of the chlorine, and the forceps, disinfected by the chlorine, may be used to plate the particles. Studies in our laboratory have shown that the treatment outlined here may be inadequate under some conditions. In commodities such as peanuts or maize where high levels of Aspergillus flavus or Penicillium species may be psent, surface disinfection may be difficult. Here 2 min immersion in 70% ethanol followed by 2 min in 0.4% chlorine is recommended. Rinsing. After the chlorine is poured off, particles may be rinsed once with sterile water. Use a 1 min treatment, with stirring, then pour the water off. Again a watch glass should cover the beaker during this period, and the sterile forceps should be used for stirring. It is not clear whether rinsing fulfils any essential function. Early direct plating regimens used agents such as mercuric chloride for disinfection, and rinsing was essential to remove such toxic materials before they penetrated the particles too deeply. However, chlorine is effectively denatured by the particles and is believed to penetrate very little. In our opinion, the rinsing step can be omitted without loss of efficacy of the treatment, with savings in time and materials, and reduced risk of recontamination from the air. Plating. After disinfection and the optional rinse, particles should be plated onto solidified agar, at the rate of 6-20 particles per plate, depending on

4.2 Enumeration Techniques

particle size. Use the disinfected forceps. Plating should be carried out immediately: keep the watch glass on the beaker if this is not possible. Incubation. Incubate plates upright, under normal circumstances for 5 days at 258C. See more detailed notes below. Examination. After incubation, examine plates visually, count the numbers of infected particles and expss results as a percentage. Differential counting of various genera is often possible. Correct choice of media, a stereomicroscope and experience will all assist in this process.

4.2.2 Dilution Plating Dilution plating is the appropriate method for mycological analysis of liquid or powdered foods. It is also suitable for grains intended for flour manufacture and other situations where total fungal contamination is relevant. Sample pparation. The two most common methods of sample pparation for dilution plating are stomaching and blending: stomaching is recommended by ICFM (King et al., 1986). The Colworth Stomacher (Sharpe and Jackson, 1972), or equivalent equipment (e.g. BagMixer1, Interscience, Saint Nom La Brete`che, France), is a very effective device for dispersing and separating fungi from finely pided materials such as flour and spices, and soft foods such as cheeses and meats, and its use is strongly recommended. Treatment time in the stomacher should be 2 min. Harder or particulate foods such as grains, nuts or dried foods, like dried vegetables, should be soaked before stomaching. Soaking times from 30 to 60 min are generally sufficient, but for extremely hard particles such as dried legumes, soaking for up to 3 h may be required. Comminution in a Waring Blender or similar machine is a suitable alternative for these types of samples and may give a more satisfactory homogenate. Blending times should not exceed 60 sec, as longer treatments may fragment mycelium into lengths too short to be viable or overheat the homogenate. The sample size used should be as large as possible. If a stomacher or BagMixer 400 is used, a sample size of 10-40 g is suitable.

21

Diluents. The recommended diluent is aqueous 0.1% peptone (Hocking et al., 2006a, p. 344), suitable for both filamentous fungi and yeasts. Saline solutions, phosphate buffer or distilled water are no longer recommended by ICFM as they may be deleterious to yeasts (Mian et al., 1997). The addition of a wetting agent such as polysorbate 80 (Tween 80) may be desirable for some products, but the natural wetting ability of the peptone is usually adequate. Special diluents may be necessary in some circumstances. If yeasts are to be enumerated from dried products or juice concentrates, the diluent should also contain 20-30% sucrose, glucose or glycerol, as the cells may be injured or be susceptible to osmotic shock. Dilution. Serial dilutions of fungi are carried out by the same procedures as those used in bacteriology, and the recommended dilution rate is 1:10 (=1+9). Fungal spores sediment more quickly than bacteria, so it is important to draw aliquots for dilution or plating as soon as possible, pferably within 1 min (Beuchat, 1992). Plating. Spad (surface) plating is recommended. When pour plates are used, fungi develop more slowly from beneath the agar surface and may be obscured by faster growing colonies from surface spores. Hence spad plating allows more uniform colony development, improves the accuracy of enumeration of the colonies and makes subsequent isolation of pure cultures easier. The optimum inoculum for surface plating is 0.1 ml. Best results will be obtained if plates are dried slightly before use. After adding the inoculum, spad it evenly over the agar surface with a sterile bent glass rod (”hockey stick”). Sterilise the rod by flaming it with ethanol before use. A plate spinner is a useful accessory. It is usually possible to enumerate plates with up to 150 colonies, but if a high proportion of rapidly growing fungi are psent, the maximum number which can be distinguished with any accuracy will be lower. Because of this restriction on maximum numbers, it may be necessary on occasion to accept counts from plates with as few as 10-15 colonies. Clearly, such limitations on numbers per plate and the overgrowth of slow colonies will result in higher counting errors than are usually achieved with bacteria or yeasts. Enumerating yeasts is less difficult. In the absence of filamentous fungi, from 30 to 300 colonies per

22

plate can be counted and errors will be comparable with those to be expected in bacterial enumeration. Incubation. The standard incubation conditions are 258C for 5 days. However, other conditions may be more suitable in some circumstances (see notes below). Reporting results. As in bacteriology, results from dilution plating are expssed as viable counts per gram of sample. Note that such results are not directly comparable with those obtained from direct plating and may not offer a direct indication of the extent of fungal growth.

4.2.3 Incubation Conditions As noted above, the standard incubation conditions specified by ICFM are 258C for 5 days (Hocking et al., 2006a). Undoubtedly, 258C is the most suitable temperature for routine work in temperate to subtropical environments. Few if any common fungi are sensitive to this temperature, even those which grow readily under refrigeration. Higher temperatures are unacceptable in the temperate zone: 308C is close to the upper limit for growth of some important Penicillium species. In tropical regions, incubation at 308C is recommended as a more realistic temperature for enumerating fungi from commodities stored at ambient temperatures. In cool temperate regions such as Europe, 228C may be a more suitable incubation temperature. When used for fungi, Petri dishes should be stored upright. The principal reason is that some common fungi can shed large numbers of spores during handling, which in an inverted dish will be transferred to the lid. Reinversion of the Petri dish for inspection or removal of the lid may liberate spores into the air or onto benches and cause serious contamination problems.

4.3 Sampling Surfaces Methods are outlined below for directly sampling the mycoflora of surfaces of commodities such as fruits, meats, cheeses, salamis and dried fish and also packaging materials, machinery and walls.

4 Methods for Isolation, Enumeration and Identification

The techniques are based on those described by Langvad (1980) for studying the fungal flora of leaves. If samples are particulate, or can be cut up, sterile forceps can be used to pss pieces of a suitable size (up to about 10 mm2) onto a suitable medium in a Petri dish. The sample is then removed, leaving an impssion, and any spores or mycelium transferred will form colonies within a few days. This technique is known as pss plating. For packaging materials such as cardboard, an alternative method is to cut a piece which will fit in a standard Petri dish. The dish is ppared by adding a sterile filter paper moistened with 10% glycerol and then placing a bent glass rod on it as a separator. After adding the sample, a thin layer of an appropriate agar medium is poured over its surface. To reduce evaporation, the dish should be sealed with Parafilm or a similar material or placed in a polyethylene bag before incubation at 258C for a few days. If contamination levels are not too heavy, the number and types of moulds psent can be effectively estimated by this method. Colonies may be subcultured for identification. For sampling walls or other surfaces, or for nondestructive sampling, impssions may be taken using adhesive tape. Carefully handled, tape coming from the roll will be virtually sterile. Press a short length of tape firmly onto the surface to be sampled, adhesive side down, then transfer it, still with the same side down, onto a suitable growth medium. After 1-2 days incubation at 258C, the tape may be removed to allow development and sporulation of colonies. Surface sampling techniques for assessment of sanitation in food production and processing areas are discussed by Evancho et al. (2001). Surfaces may be sampled by using sterile swabs or by agar contact methods. The swab method involves rubbing a moistened sterile cotton swab over the test surface and placing the swab in a dilution bottle to be subsequently diluted and plated on appropriate media. Agar contact plates, also known as RODAC (replicate organism direct agar contact) plates, are restricted to use on smooth or semi-smooth flat surfaces. An alternative to agar contact plates is the agar slice technique, where agar is filled into a syringe-like apparatus or into an artificial sausage casing. The solidified agar can be pushed out onto

4.5 Isolation Techniques

the surface to be sampled, then the portion making contact sliced off with a sterile scalpel or wire and placed in a Petri dish for incubation.

4.4 Air Sampling Air sampling in the food processing environment is discussed by Evancho et al. (2001). The simplest method of air sampling is by sedimentation or settle plates. A Petri dish containing an appropriate agar medium is exposed to the atmosphere for a fixed period of time (usually 15-60 min), then closed and incubated at 258C (Samson et al., 2004a). This method can be useful in food production areas, as it gives a direct indication of the number and types of fungi likely to come into contact with exposed product. However, the settle plate technique lacks pcision, and volumetric air sampling is a much more reliable indicator of air quality. A number of air sampling devices are commercially available. Of these, the Anderson sampler (either the two-stage or six-stage model; Anderson Instruments Inc, Atlanta, GA, USA) is probably the best, giving accurate and consistent results (Buttner and Stetzenbach, 1993). However, the Anderson sampler is expensive and requires mains power or a large battery for operation. In factory situations, the dry cell battery-operated Biotest

23

Hycon RCS and RCS High Flow centrifugal air samplers (Biotest, Solihull, UK) and the MAS-100 air sampler (Merck, Darmstadt, Germany) are more convenient, as they are small and readily portable. The Biotest RCS Plus sampler was reported to give comparable results to larger and more sophisticated machines, but its sampling efficiency gradually decreased for particle sizes below 4 mm (Benbough et al., 1993).

4.5 Isolation Techniques The term ”isolation” is used here in its strict sense: the pparation of a pure culture, free from any contamination and ready for identification.

4.5.1 Yeasts Streaking techniques commonly used for bacterial purification are equally suitable for the isolation of yeasts (Fig. 4.1). A method widely used by yeast specialists is to disperse a portion of a colony in 2- 3 ml of sterile water, then streak a single loop of this suspension over the whole surface of a plate, moving the loop slowly down from top to bottom while simultaneously moving it rapidly across the plate

Fig. 4.1 Petri dishes of yeasts growing on malt extract agar, showing a streaking method suitable for producing isolated colonies

24

from side to side. After suitable incubation, wellseparated single colonies should appear in the lower half of the plate (Fig. 4.1). If all of these single colonies appear to be of similar size and appearance (taking into account the effect of crowding), the culture may be judged to be pure. Microscopic checks of some single colonies are also desirable. Disperse a needle point of cells from a colony in a drop of water, add a cover slip and examine by bright field illumination at about 400. Cell outlines will be clearly visible. Note that, unlike those of bacteria, yeast cell sizes often vary considerably in a pure pparation. Purity is indicated not so much by uniformity of cell size within a pparation as by similarity in microscopic cell appearance from colony to colony. When a culture is considered to be pure, streak it onto an appropriate slant.

4.5.2 Moulds Streaking techniques are ineffective for filamentous fungi and are not recommended. Isolation depends on picking a small sample of hyphae or spores – judged to be pure by eye, by hand lens or pferably under the stereomicroscope – and placing this sample on a fresh plate as a point inoculum. Purity is subsequently judged by uniformity in appearance of the colony which forms after incubation. The appearance of a mixed culture depends on the growth rates of the fungi psent. If rates are perse, a mixed culture is often indicated by a clump of dense hyphae at the inoculum point, surrounded by looser wefts of spading hyphae. With fungi of approximately equal growth rates, mixtures are often indicated by colonies with sectoring growth: sectors will show differences in mycelial, spore or reverse colours or in radial growth rates. The simplest starting place for isolating fungi is an enumeration plate with well-separated colonies. Use a needle of platinum or nichrome, pferably cut to a chisel point with a pair of pliers, or a steel sewing needle. Sterilise it by heating, then plunge the tip into cold agar and leave until cool – with nichrome or steel this will require several seconds. With the tip of the cold, wet needle pick off a few spores or a tuft of mycelium – just enough to be

4 Methods for Isolation, Enumeration and Identification

4.6 Choosing a Suitable Medium

25

require free access to oxygen for typical growth and sporulation. Oxygen starvation during growth will at best lead to retarded sporulation or at worst death of the culture. Long-term pservation of fungi is dealt with later in this chapter.

4.6 Choosing a Suitable Medium Food laboratories often rely on a single all-purpose medium to produce a ”yeast and mould” count in everything from raw material to finished product. But just as the food bacteriologist uses selective media for particular purposes, so too food mycologists are developing a range of media suited to specific applications. It is plainly unrealistic to expect a single medium to answer all questions about mould and yeast contamination in all foods. The fungi that spoil meats or fresh vegetables are not the same as those that grow on dried fish. Although often used for this purpose, very dilute media such as potato dextrose agar are of little or no value for enumerating fungi from dried foods.

The most important pision in types of enumeration media lies between those suitable for high water activity foods, such as eggs, meat, vegetables and dairy products, and those suited to the enumeration of fungi in dried foods. The most suitable media for dried foods depend on the type of food, the major categories being foods low in soluble solids such as cereals, high sugar foods such as confectionery and dried fruit, and salt foods. A second consideration lies in whether the primary interest is in moulds, or yeasts, or both; and a third concerns the psence or absence of pservatives. Finally, media are available for specific mycotoxigenic fungi, notably Aspergillus flavus and related species and Penicillium verrucosum plus P. viridicatum. An overview of media considered most suitable for particular purposes is given in Table 4.1. The table is derived from Pitt and Hocking (1997), together with recent recommendations from ICFM (Samson et al., 1992; Hocking et al., 2006a).

Table 4.1 Recommended media for fungal detection, enumeration and isolationa Type of food

Selecting for

Medium

Remarks

Fresh foods: milk and milk products, fruit, cheese, sea foods

Moulds Yeasts General

DRBC TGY, MEA, OGY DRBC

Blend (where necessary) and dilution plate

Freshly harvested grains, nuts

General Dematiaceous Hyphomycetes Fusarium Yeasts

DRBC DRBC, CZID CZID TGY, MEA, OGY

Direct plate Direct plate Direct plate Dilution plate

Fruit juices, fresh

Yeasts

TGY, MEA, OGY

Dilution plate

Fruit juices, pserved

Preservative resistant yeasts

TGYA, malt acetic agar

Dilution plate

Fruit juices, to be pasteurised, or pasteurised products

Heat resistant moulds

PDA, MEA

Special protocol

Fruit juice concentrates

Xerophilic yeasts

MY50G

Special diluents

Dried foods in general

General

DG18

Direct plate

Stored cereals, nuts

General Dematiaceous Hyphomycetes Fusarium

DG18 DRBC, CZID CZID

Direct plate Direct plate Direct plate

26

4 Methods for Isolation, Enumeration and Identification

Table 4.1 Recommended media for fungal detection, enumeration and isolationa (continued) Type of food

Selecting for

Medium

Remarks

Grain for milling into flour

General

DG18

Stomach or blend and dilution plate

Dried fruit, confectionery, chocolate, etc.

Xerophilic moulds and yeasts

MY50G

Direct plate

Fastidious xerophiles – in psence of Eurotium spp.

MY50G MY70GF

Direct plate Direct plate

General

DG18

Halophilic xerophiles

MY5-12, MY10-12

Direct plate or pss plate Direct plate or pss plate

Fungi producing aflatoxins

AFPA

Direct or dilution plate

DRYS

Direct or dilution plate

Salt foods, e.g. salt fish

General

General Fungi producing ochratoxins a For medium acronyms, see Section 4.6.

4.6.1 General Purpose Enumeration Media To be effective, a general purpose enumeration medium must fulfil several requirements (Pitt, 1986). As these are sometimes overlooked, they are listed here:

to inhibit bacterial growth completely, without

affecting growth of foodborne fungi (filamentous or yeasts); to be nutritionally adequate and support the growth of fastidious fungi; to suppss the growth of rapidly spading fungi, especially the Mucorales, but not to pvent their growth entirely, so they too can be enumerated; to slow radial growth of all fungi, to permit counting of a reasonable number of colonies per plate, without inhibiting spore germination; to promote growth of relevant fungi; and to suppss growth of soil fungi or others generally irrelevant in food spoilage.

Fulfilling the above requirements necessitates the use of potent inhibitory compounds, and there is sometimes a fine line between inhibition of undesirable microorganisms and suppssion of growth of those being sought. Modern fungal enumeration media rely on the use of antibiotics at neutral pH

for the inhibition of bacteria. Such media allow better recovery of moribund and sensitive fungi than the acidified media commonly used in the past. For many years rose bengal has been added to media to slow colony spad, while more recently 2,6-dichloro-4-nitroaniline (dichloran) has been added to inhibit rapidly spading moulds. Many common spoilage fungi, Aspergillus and Penicillium species in particular, develop better on media with adequate nutrients. Low nutrient media of very high aw, such as potato dextrose agar, have lost favour because they are selective against some species in these genera. The media described below are considered to be the most satisfactory general purpose enumeration media available at this time (Hocking et al., 2006a). Formulations are given in the Media Appendix. Dichloran rose bengal chloramphenicol (DRBC) agar. DRBC (King et al., 1979, Pitt and Hocking, 1997) is recommended for both moulds and yeasts. It is particularly suited to fresh and high aw foods (Hocking et al., 1992). This medium contains both rose bengal (25 mg/kg) and dichloran (2 mg/kg), which restrict colony spading without affecting spore germination unduly. Compact colonies allow crowded plates to be counted more accurately. This combination of inhibitors also effectively restricts the rampant growth of most of the common mucoraceous fungi such as Rhizopus and

4.6 Choosing a Suitable Medium

27

Fig. 4.2 Petri dishes of (a) DRBC and (b) RBC showing effective control of Rhizopus growth by rose bengal and dichloran in DRBC

Mucor (Fig. 4.2), although it does not completely control some other troublesome genera such as Trichoderma. In routine use, it is recommended that DRBC plates be incubated away from light at 258C for 5 days. Dichloran 18% glycerol (DG18) agar. Hocking and Pitt (1980) developed DG18 to be selective for xerophilic fungi from low moisture foods such as stored grains, nuts, flour and spices. DG18 was designed for enumeration of a range of nonfastidious xerophilic fungi and yeasts. However, it has been shown since that it supports growth of the common Aspergillus, Penicillium and Fusarium species, as well as most yeasts, and many other common foodborne fungi. DG18 can now be described as a general purpose medium with emphasis on enumeration of fungi from dried foods. DG18 is also recognised to be a very useful medium for enumeration of airborne fungi (Wu et al., 2000; Samson et al., 2004a). It is an effective inhibitor of Mucoraceous fungi, and bacteria are totally suppssed. However, growth of Eurotium species (pviously known as ”the Aspergillus glaucus group”) is still somewhat too rapid, and colonies may have diffuse margins. The addition of detergent to DG18 has been reported to be an improvement in this respect (Beuchat and de Daza, 1992). Although DG18 is a satisfactory isolation medium for Eurotium species, it is not suitable for their

identification. Eurotium species are usually identified on Czapek yeast extract agar with 20% sucrose (CY20S), described later in this chapter. Other general purpose media. Under circumstances where rapidly spading moulds do not cause problems, two alternative general purpose enumeration media are satisfactory. These are rose bengal chloramphenicol agar (RBC; Jarvis, 1973), from which DRBC was developed, and oxytetracycline glucose yeast extract agar (OGY; Mossel et al., 1970). OGY has been found to be very suitable for yeasts in the absence of moulds (Andrews, 1992a). Most of the media discussed above are available in ready to use dehydrated form from media suppliers such as Oxoid, Difco, BBL, etc.

4.6.2 Selective Isolation Media Although considerable progress has been made in the past 20 years, the formulation of selective media for foodborne fungi still requires a great deal of research. The availability of effective media can greatly simplify the isolation and identification of significant food spoilage and mycotoxigenic fungi. Most attention has been paid so far to the requirements of xerophilic fungi because of their failure to develop on standard high aw media. For mycotoxigenic fungi, satisfactory media exist only for

28

4 Methods for Isolation, Enumeration and Identification

Coconut Cream agar (CCA) to detect aflatoxin production in Aspergillus flavus and A. parasiticus. CCA can be made using any brand of commercial canned coconut cream (available from Asian food stores in many places). Dilute 50:50 with water, add agar (1.5%) and autoclave. Inoculate solidified plates with up to four colonies (picked with a wet needle) and incubate at 308C for 5-7 days. Examine, reverse upmost, under long wave length UV light. Colonies producing aflatoxins will fluoresce bluish white or white, especially in the centres. Ignore fluorescence from the Petri dish itself. Use an uninoculated coconut cream agar plate as a control (Dyer and McCammon, 1994). Plates inoculated with known nontoxigenic and toxigenic strains are also useful controls. A. parasiticus isolates almost always produce aflatoxins. Media for fungi producing ochratoxin A. Although ochratoxin A was first described from Aspergillus ochraceus, recent molecular studies indicate that A. westerdijkiae is the major ochratoxin A producing species in Aspergillus Section Circumdati (Frisvad et al., 2004). Aspergillus carbonarius is also an important source of ochratoxin A, particularly in grape products (Abarca et al., 2004; Leong et al., 2006a and references therein). However, there are no selective or indicative media for these fungi. When DG18 is used as the isolation medium, selection of colonies with light brown (ochre) or black sporulation is a good starting point for detecting A. westerdjikiae or A. carbonarius respectively. Coconut cream agar (Dyer and McCammon, 1994) can also be used to screen for ochratoxin production in A. carbonarius and species in Aspergillus Section Circumdati (Heenan et al., 1998). Plates are best incubated at 258C rather than 308C for potentially ochratoxigenic species of Aspergillus. Penicillium verrucosum is the most important ochratoxin a producer in the genus Penicillium (Pitt, 1987), although P. nordicum can also produce this mycotoxin (Frisvad and Samson, 2004). Frisvad (1983) developed Dichloran rose bengal yeast extract sucrose agar (DRYS) for selective enumeration of P. verrucosum. P. nordicum, P. viridicatum and P. aurantiogriseum are also selected by DRYS. P. verrucosum and P. nordicum produce ochratoxin A, P. verrucosum also produces citrinin, while the latter two species produce xanthomegnin and viomellein. According to Frisvad (1983), P. verrucosum colonies on DRYS have a violet brown reverse, and

4.6 Choosing a Suitable Medium

the latter two species produce yellow colonies with a yellow reverse. The incubation regimen recommended by Frisvad (1983) is 7-8 days at 208C. Subsequently, Frisvad et al. (1992) developed dichloran yeast extract sucrose 18% glycerol agar (DYSG) on which P. verrucosum produces a red brown reverse. Because of its lower aw, DYSG inhibits rapidly growing fungi such as Rhizopus and Mucor more effectively than DRYES. In a study of 76 samples of wheat, rye and barley, Lund and Frisvad (2003) found that DYSG was more effective than DRYES for screening grain samples for the psence of P. verrucosum and potential ochratoxin A contamination. Media for Fusarium species. Dichloran chloramphenicol peptone agar (DCPA; Andrews and Pitt, 1986) can be used to isolate Fusarium species from grains and other substrates. The medium was developed from Nash-Snyder medium (Nash and Snyder, 1962) a medium for enumeration of Fusarium species from soils. DCPA uses a low level of dichloran as a substitute for the high level of pentachloronitrobenzene (PCNB), and chloramphenicol rather than the antibiotic mixture used by Nash and Snyder (1962). When Fusarium species are dominant, DCPA is effective for their isolation from grains, animal feeds and soil. However, DCPA has been found to be less effective in mixed populations. DCPA is a very useful medium for the identification of Fusarium species because it often induces the abundant formation of macroconidia (Hocking and Andrews, 1987). A much more stringent medium, effective for isolation of most Fusarium species occurring in foods, is Czapek iprodione dichloran agar (CZID; Abildgren et al., 1987). As well as dichloran, this medium contains the fungicide iprodione. CZID is highly selective for Fusarium species, and is probably the best medium for Fusarium enumeration and isolation. CZID is suitable for isolation of Fusarium species by direct plating of surface disinfected grains and other commodities, or dilution plating of homogeneous samples such as flour. Some questions remain concerning whether CZID may be too selective and not support growth of all foodborne Fusarium species. However, the common species all grow well. Castella´ et al. (1997) developed a Fusarium selective medium (MGA 2.5) using 2.5 mg/L malachite green as the selective agent. They reported that MGA 2.5 was more selective than Nash-Snyder medium as it did not allow development of colonies of other fungal genera.

29

Bragulat et al. (2004) compared the efficacy of MGA 2.5 medium with DCPA, CZID and several other Fusarium media, using pure cultures of twelve Fusarium species commonly found in foods as well as naturally contaminated samples. They reported that there was no statistically significant difference in colony counts of the Fusarium spp. tested, but that colonies on MGA 2.5 were smaller than the other media. MGA 2.5 did not allow growth of other fungi such as Zygomycetes and yeasts from naturally contaminated samples, thus providing better selectivity than the other media. Media for dematiaceous Hyphomycetes. Although designed primarily for Fusarium isolation, CZID has been found in our laboratory to be very useful for isolating dematiaceous Hyphomycetes, provided that iprodione is added at half the usual concentration, as full strength iprodione tends to restrict colony diameters too severely. Alternaria, Bipolaris, Curvularia, Stemphylium and Ulocladium species will grow and sporulate well when incubated for 5 days at 258C with a 12 h photoperiod. Drechslera species will grow but will not sporulate on this medium. DRBC is also of value for isolating these fungi, but some species do not sporulate readily on it. DCPA, originally developed for isolation of Fusarium, may also be used as an isolation medium for the dematiaceous Hyphomycete genera mentioned above. Alternaria, Curvularia and related genera usually grow rapidly and sporulate well on DCPA. However, fungi should not be maintained or stored on DCPA for more than two weeks, as ammonia is produced by aging cultures. Media for xerophilic fungi. Xerophilic fungi are of great importance in food spoilage, and hence media and techniques for their enumeration and isolation have received much attention. Xerophiles range from those which grow readily on normal media and which are only marginally xerophilic, such as many Aspergillus and Penicillium species, to those, such as Xeromyces bisporus, which will not grow at all on normal (high aw) media. It is not surprising that no single medium is suitable for quantitative estimation of all xerophilic fungi found causing food spoilage. As noted earlier, DG18 was developed as a general medium for xerophiles, and remains the medium of choice for this purpose. DG18 should be used in any general examination of the mycoflora of dried foods. Media for extreme xerophiles. Fungi discussed here include Xeromyces bisporus, xerophilic

30

4 Methods for Isolation, Enumeration and Identification

The most satisfactory medium is malt extract yeast extract 70% glucose fructose agar (MY70GF). MY70GF is of similar composition to MY50G, except that it contains equal parts of glucose and fructose to pvent crystallisation of the medium at the concentration used (70% w/w). It is made in a similar manner to MY50G. Growth of even the extreme xerophiles on MY70GF is extremely slow, and plates should be incubated for at least 4 weeks at 258C. Once growth is apparent, pick off small portions of colonies and transfer them to MY50G, to allow more rapid growth and sporulation. Media for halophilic xerophiles. Some xerophilic fungi from salted foods such as salt fish grow more rapidly on media containing NaCl and hence are correctly termed halophilic xerophiles. Malt extract yeast extract 5% salt 12% glucose agar (MY5-12) and malt extract yeast extract 10% salt 12% glucose agar (MY10-12) are suitable for these fungi. Techniques for enumeration and isolation are similar to those described above for other extreme xerophiles.

4.6.3 Techniques for Yeasts The simplest enumeration and growth medium for most food spoilage yeasts is malt extract agar (MEA). Although originally introduced as a growth medium for moulds, its rich nutritional status makes it very suitable for yeasts, and its relatively low pH (usually near 5.0) reduces the possibility of bacterial contamination. More recently, tryptone glucose yeast extract agar (TGY) has been recommended for enumeration of yeasts (Hocking et al., 1992, 2006a). Due to a higher glucose concentration (10%) and higher pH (5.5-6.0), this medium is more effective in recovery of stressed yeast cells, and colony development is usually faster than on MEA. However, its higher pH means that an antibiotic should be incorporated for enumeration of yeasts from food samples which may contain bacteria. Recommended antibiotics are chloramphenicol or oxytetracycline at the concentrations used in DRBC and OGY. Both MEA and TGY are suitable for enumeration of yeasts in products such as fruit juices, fruit purees and yoghurt, where moulds are usually psent only in low numbers.

4.6 Choosing a Suitable Medium

31

and replace it with sterile water. Leave the cap loose, incubate at room temperature or 308C and watch for evidence of fermentation. Shaking the container daily will help to detect gases resulting from fermentation. Classical enrichment techniques used in bacteriology can also be used for yeasts. TGY broth has been used very successfully in our laboratory for enrichment of low numbers of yeasts in liquid products. Add 10 ml of product to 90 ml TGY broth and incubate at 258C for 3-4 days, or 308C for 2-3 days. Look for signs of fermentation and streak out onto TGY agar. Detection of pservative resistant yeasts. A few species of yeasts are able to grow in products containing pservatives such as sorbic, benzoic and acetic acids or sulphur dioxide. The most important of these is Zygosaccharomyces bailii. The simplest and most effective way to screen for pservative resistant yeasts is to spad or streak product onto plates of malt acetic agar (MAA), which is MEA with 0.5% acetic acid added (Pitt and Richardson, 1973) or TGY with 0.5% acetic acid added (TGYA, Hocking, 1996). MAA and TGYA are made by adding glacial (16 N) acetic acid to melted and tempered basal medium to give a final concentration of 0.5%. Mix and pour immediately. These media cannot be held molten for long periods or remelted because of their low pH (approximately 3.2 and 3.8 for MAA and TGYA, respectively). The acetic acid does not need sterilisation before use. MAA and TGYA are suitable media for monitoring raw materials, process lines and products containing pservatives for resistant yeasts. They are also effective for testing pviously isolated yeasts for pservative resistance. Erickson (1993) developed a selective medium for Zygosaccharomyces bailii. Zygosaccharomyces bailii medium (ZBM) is based on Sabouraud dextrose agar amended with fructose, NaCl, tryptone, yeast extract and trypan blue dye, then acetic acid (0.5%) and potassium sorbate (0.01%) are added to make the medium selective. It is designed as a plating medium for detection of Z. bailii in acidified ingredients in conjunction with hydrophobic grid membrane filtration (Erickson, 1993). When compared with MAA and TGYA in an interlaboratory study, ZBM was found to be highly selective for Zygosaccharomyces bailii, to the exclusion of other important pservative resistant yeasts such as Schizosaccharomyces pombe and Pichia

32

membranaefaciens. In addition, recovery of Z. bailii cells sublethally injured by lyophilisation was significantly lower on ZBM than on TGYA or MAA (Hocking, 1996). Its high selectivity and complex formulation make ZBM unsuitable for routine laboratory use as a medium for detection of pservative resistant yeasts. Enrichment of pservative resistant yeasts. A technique capable of detecting yeast numbers as low as 1 cfu/ml within 4 days has been developed in our laboratory (Hocking et al., 1996). This method is particularly suitable for the detection of Zygosaccharomyces bailii in raw materials or finished product, but can also be used for the detection of Schizosaccharomyces pombe, Pichia membranaefaciens and other species of pservative resistant yeasts. The method involves a 2to 3-day enrichment step followed by a plating step with a further 2 days of incubation. Triplicate 20 ml tryptone glucose yeast extract (TGY) broths containing 0.5% acetic acid (TGYA) are each inoculated with 1 g or 1 ml of product and the broths incubated at 308C. After incubation for 48 and 72 h, 0.1 ml from each broth is spad plated onto TGY agar containing 0.5% acetic acid and the plates incubated at 308C. The detection time of the method is shortened by incubating broths and plates at 308C rather than the traditional temperature of 258C, as the optimum growth temperature for Zygosaccharomyces bailii is 30-328C (Jermini and Schmidt-Lorenz, 1987b). The sensitivity of the method is greatly increased by using triplicate broths instead of single or duplicate broths and by spad plating 0.1 ml from each broth instead of streaking a loopful onto TGYA agar. This method has been used to detect low numbers of cells of Zygosaccharomyces bailii in experimentally inoculated cordial syrup, mayonnaise, salad dressing and barbecue sauce and other pservative-resistant yeasts such as Schizosaccharomyces pombe, Pichia membranaefaciens and some pservative resistant strains of Saccharomyces cerevisiae. Yeasts of intermediate pservative resistance (e.g. Debaryomyces hansenii, Candida krusei and Torulaspora delbrueckii) can also be detected by this method. Better recoveries were obtained using TGY than a malt extract agar and broth system, possibly due to the fact that the pH of TGY broth + 0.5% acetic acid is 3.8, compared with pH 3.2 for MEA + 0.5% acetic acid, and there is a lower concentration of glucose (2%) in the ME

4 Methods for Isolation, Enumeration and Identification

system compared with 10% in TGY. Yeasts which are unable to grow in the psence of acetic acid or other weak acid pservatives (sorbic or benzoic acids and their salts) are not detected by this method.

4.6.4 Techniques for Heat-Resistant Fungi Heat resistant spoilage fungi, such as Byssochlamys, Talaromyces, Neosartorya and Eupenicillium species can be selectively isolated from fruit juices, pulps and concentrates by laboratory pasteurisation using various methods (Beuchat and Rice, 1979; Hocking and Pitt, 1984; Beuchat and Pitt, 2001; Houbraken and Samson, 2006). Three methods are described here: a plating method based on that of Murdock and Hatcher (1978), a direct incubation method and a filtration method for liquid samples such as liquid sugar. Plating method. If the sample to be tested is more concentrated than 358Brix, it should first be diluted 1:1 with 0.1% peptone or similar diluent. For very acid juices such as passionfruit, normally about pH 2.0, the pH should be adjusted to 3.5-4.0. Two 50 ml samples are taken for examination. Erlenmeyer flasks (250 ml) or polyethylene Stomacher bags may be used as heat penetration into these containers will be rapid. If using Stomacher bags, the tops should be heat sealed. If a heat sealer is not available, the tops may be folded over and secured with a clip and should not be fully immersed. The two samples are heated in a closed water bath at 808C for 30 min, then rapidly cooled. Each 50 ml sample is then mixed with an equal volume of double strength MEA distributed over four 150 mm Petri dishes. The Petri dishes are loosely sealed in a plastic bag to pvent drying and incubated at 308C for up to 30 days. Plates are examined weekly for growth. Most moulds will produce visible colonies within 10 days, but incubation for up to 30 days will allow for the possible psence of badly heat damaged spores, which may germinate very slowly. This long incubation time also allows most moulds to mature and sporulate, aiding their identification. The main problem associated with this technique is the possibility of aerial contamination of the plates with common mould spores, which will give false positive results. The growth of green Penicillium

4.6 Choosing a Suitable Medium

colonies, or colonies of common Aspergillus species such as A. flavus and A. niger, is a clear indication of contamination as these fungi are not heat resistant. To minimise this problem, plates should be poured in clean, still air or a Class 2 biohazard cabinet if possible. If a product contains large numbers of heat resistant bacterial spores (e.g. Bacillus species), antibiotics can be added to the agar. The addition of chloramphenicol (100 mg/l of medium) will pvent the growth of these bacteria. Direct incubation method. A more direct method used for screening fruit pulps and other semisolid materials avoids the problems of aerial contamination. Place approximately 30 ml of pulp in each of three or more flat bottles such as 100 ml medicine flats. Heat the bottles in the upright position for 30 min at 808C and cool, as described pviously. The bottles of pulp can then be incubated directly, without opening and without the addition of agar. They should be incubated flat, allowing as large a surface area as possible, for up to 30 days at 308C. Any mould colonies which develop will need to be subcultured onto a suitable medium for identification. If containers such as Roux bottles are available, larger samples can be examined by this technique, but heating times must be increased. Bottle contents should reach at least 758C for 20 min when checked by a thermometer suspended near the centre of the pulp. For further details of the above methods see Hocking and Pitt (1984) or Beuchat and Pitt (2001). Filtration method for liquid sugars. This method permits the detection of very low numbers of cells in clear liquids such as liquid sugar. Sample size should be at least 100 g, taken after vigorously shaking the container from which the sample is drawn. Add 100 ml diluent (0.1% aqueous peptone) to 2 50 g samples and mix well to dissolve. Filter both samples sequentially through the same sterile 0.45 mm membrane filter. After both samples have passed through the filter, rinse the interior of the funnel with 3 20- 30 ml volumes of sterile diluent. Remove the filter from the filter holder using sterile forceps and place it in a sterile bottle or Stomacher bag. Add 10 ml diluent to the bottle or bag containing the filter and place in a water bath at 758C for 30 min. Ensure the sample is submerged in the water bath (weigh down if necessary). Cool rapidly to room temperature, shake well, then pide the 10 ml of diluent between three Petri dishes. Add a generous portion of MEA with

33

antibiotics to each plate, mix the agar and sample well, then let the plates solidify. Incubate at 308C for up to 30 days, examining weekly. Count colonies and report count per 100 g. This method was developed by BCN Research Laboratories, Rockford, Tennessee, USA.

4.6.5 Other Plating Techniques Three other techniques which were developed for counting bacteria have been applied to fungal enumeration. Spiral plate count. Zipkes et al. (1981) evaluated the application of the spiral plate procedure to the enumeration of yeasts and moulds. They compared this procedure with the traditional pour plate and streak plate methods and found that spiral plating gave a higher overall recovery and lower replicate plating errors than the other two methods. The medium they used was potato dextrose agar, but the technique should be no less efficient using the media recommended here. Automation of spiral plate counting was studied by Manninen et al. (1991). They compared counts of pure bacterial, yeast and mould cultures using standard plating methods and spiral plate counts determined both manually and with a laser colony detector. They concluded that counts were not significantly different except where large colonies (10-15 mm diameter) of Rhizopus oligosporus were enumerated. Other Rhizopus and Mucor species are also likely to interfere with this method unless a suitable plating medium (such as DRBC) is used. Alonso-Calleja et al. (2002) found that the spiral plate technique compared well with standard plate counting on OGYE agar for enumeration of yeasts and moulds in goat’s milk cheeses; however, Garcı´ a-Armesto et al. (2002) found the method unsuitable for enumeration of yeasts in raw ewe’s milk. Hydrophobic grid membrane filters. Membrane filters overprinted with a square hydrophobic grid have been developed for rapid enumeration of bacteria. The hydrophobic grid membrane filter TM (HGMF) system is marketed as ISO-Grid by Neogen (Lansing, MI, USA). The HGMF ”count” is determined by a most probable number (MPN) calculation. Brodsky et al. (1982) applied the HGMF technique to counting yeasts and moulds in foods. They compared it with spad plating on

34

4 Methods for Isolation, Enumeration and Identification

4.7 Estimation of Fungal Biomass A deficiency in all of the enumeration techniques which rely on culturing fungi is that the result is at best poorly correlated with growth or biomass. Biomass is usually regarded as the fundamental measure of fungal growth in biotechnology, but it is not easy to quantify under the conditions existing in foods. Mycelial dry weight is most commonly used as a biomass estimate, but its relationship to mycelial wet weight and to metabolism varies widely in foods, due to the great influence of aw on both of the latter parameters. Fungi growing at reduced aw can be expected to be more dense than at high aw due to increased concentrations of internal solutes, though this is exceptionally difficult to measure experimentally. The question of a satisfactory fundamental measure of fungal biomass remains unanswered. Despite these basic problems, several chemical and biochemical techniques are available to estimate the extent of fungal growth in commodities. These techniques rely either on some unique component of the fungus that is not found in other microorganisms or foods or on immunological or molecular techniques. Some are still in the developmental phase: the most important ones are described briefly here.

4.7.1 Chitin Chitin is a polymer of N-acetyl-D-glucosamine and is a major constituent of the walls of fungal spores and mycelium. It also occurs in the exoskeleton of insects but is not psent in bacteria or in foods. Hence the chitin content of a food or raw material can provide an estimate of fungal contamination. Chitin is most effectively assayed by the method of Ride and Drysdale (1972). Alkaline hydrolysis of the food sample at 1308C causes partial depolymerisation of chitin to produce chitosan. Treatment with nitrous acid then causes partial solubilisation and deamination of glucosamine residues to produce 2,5-anhydromannose, which is estimated colorimetrically using 3-methyl-2-benzothiazolone hydrazone hydrochloride as the principal reagent. Alkaline hydrolysis is more readily accomplished at

4.7 Estimation of Fungal Biomass

1218C in an autoclave (Jarvis, 1977). Improved assay sensitivity was achieved by derivatisation of glucosamine and other products with o-phthalaldehyde, separation by high performance liquid chromatography and detection of fluorescent compounds with a spectrofluorimeter (Lin and Cousin, 1985). Ekblad and Nasholm (1996) also described an HPLC method which measured fluorescence of a 9-fluorenylmethylchloroformate derivative of glucosamine. The chitin assay remains rather complex and slow, usually requiring about 5 h. A number of studies have indicated that the chitin assay is a valuable technique for estimating the extent of fungal invasion in foods such as maize and soybeans (Donald and Mirocha, 1977), wheat (Nandi, 1978) and barley (Whipps and Lewis, 1980) to estimate mycorrhizal fungi and fungal pathogens in plant material and soil (Ekblad and Nasholm, 1996; Ekblad et al., 1998; Penman et al., 2000; Singh, 2005) and measure wood rotting fungi (Nilsson and Bjurman, 1998). Particular attention has been paid to the possibility of developing the chitin assay as a replacement for the Howard mould count for tomato products (Jarvis, 1977; Bishop et al., 1982; Cousin et al., 1984). The chitin assay has some shortcomings and has been severely criticised by some authors (e.g. Sharma et al., 1977). The relationship between dry weight and chitin content varies at least twofold for different food spoilage fungi (Cousin et al., 1984; Lin and Cousin, 1985; Cousin, 1996). Some foods contain naturally occurring amino sugars such as glucosamine and galactosamine, which should be removed by acetone extraction prior to hydrolysis (Whipps and Lewis, 1980). Products from rot-free tomatoes give positive glucosamine assays even after acetone extraction (Cousin et al., 1984) and chitin content does not increase proportionally with fungal growth (Sharma et al., 1977). Insect contamination of grain samples has been reported to produce grossly misleading results (Sharma et al., 1977), but the psence of fruit flies in tomatobased products was less serious (Lin and Cousin, 1985). Materials such as stored grains frequently contain insect fragments and need to be checked before chitin assays are attempted. Because of these difficulties, the use of chitin as a chemical assay for fungi in foods has largely been superseded by the ergosterol assay.

35

36

mycelial mass, and varied with substrate, aeration and growth phase. The ergosterol content was low during the rapid growth phase but tended to increase, at times sharply, as growth slowed. Taniwaki (1995) demonstrated that in fungi growing in atmospheres low in O2 and with elevated CO2 levels, the ergosterol content of hyphae was significantly reduced. In atmospheres containing 60% CO2, ergosterol content per unit of hyphal length was up to six times less than in air. Growth medium also affected ergosterol concentrations: on average, seven foodborne fungal species grown on PDA produced more than twice as much ergosterol per unit hyphal length as when grown on CYA (Taniwaki, 1995; Taniwaki et al., 2006). However, there was a reasonable correlation between ergosterol and mycelium dry weight for seven of the eight species tested. Eurotium chevalieri was the exception: this species produced little ergosterol and appeared to produce several other sterols (Taniwaki et al., 2006). Marı´ n et al. (2005) examined 16 species of food spoilage fungi and concluded that ergosterol content and colony diameters were better correlated to fungal biomass than fungal counts were. Marı´ n et al. (2008) showed that, for 14 common food spoilage fungi, correlation coefficients between ergosterol and colony diameters were sufficiently significant over a range of aw values (0.95-0.85), pHs (5-7) and potassium sorbate concentrations (0.5-1.5%) for both parameters to be useful in growth modelling. Quantifying ergosterol production in foods has proved more difficult. Seitz et al. (1977) showed a good correlation between damage in rice grains and their ergosterol content, between ergosterol in wheat and rainfall during the growing season and between ergosterol content and fungal invasion in several sorghum hybrids. Matcham et al. (1985) reported good correlations between linear extension of Agaricus bisporus grown on rice grains and chitin, ergosterol and laccase production. Ergosterol content correlated with colony counts of fungi on wheat grains at 0.95 aw but not at 0.85 aw (Tothill et al., 1992). Using a stereomicroscope for visual examination, they concluded that sound grain contained up to 6 mg/g ergosterol, microscopically mouldy grain 7.5-10 mg/g and visibly mouldy grain more than 10 mg/g ergosterol. From studies on ergosterol levels, colony counts and mould growth in a variety of grain samples, Schnu¨rer and Jonsson (1992) concluded that ergosterol correlated with colony counts

4 Methods for Isolation, Enumeration and Identification

better on DG18 (r = 0.77) than on MEA (r = 0.69). Ergosterol levels of food grade wheat ranged from 2.4 to 2.8 mg/g dry weight, samples from field trials (of unspecified quality) from 3.0 to 5.6 mg/g and feed grains from 8 to 15 mg/g dry weight. After an extensive survey of ergosterol levels in Danish crops, Hansen and Pedersen (1991) concluded that the normal levels of ergosterol in barley were 7.6 – 2.8, wheat for bread making 5.0 – 1.5, rye for bread making 6.8 – 2.2, peas 2.2 – 2.7 and rapeseed 2.4 – 1.3 mg/g dry weight. Ochratoxin A in barley correlated well with ergosterol content and reached significant levels when ergosterol increased to 25 mg/g dry weight. However, aflatoxin B1 became detectable in cottonseed meal when ergosterol reached only 4 mg/g. ”Burned” rapeseed, a measure of quality, became significant when ergosterol reached 1.4 mg/g dry weight. Lamper et al. (2000) found that ergosterol content correlated well (r = 0.87) with deoxynivalenol levels in wheat inoculated with Fusarium graminearum or F. culmorum. Moraes et al. (2003) found that there was good correlation between mould counts and ergosterol content of Brazilian maize (r = 0.94) but a poor correlation (r = 0.4) between ergosterol and aflatoxin content. Pietri et al. (2004) found significant correlation between ergosterol content of Italian maize and the major mycotoxins, fumonisin B1 (1995 crop) or zearalenone and deoxynivalenol (1996 crop). Ergosterol content correlated strongly with fat acidity values and germination ability of stored canola (Pronyk et al., 2006). These authors also noted that Penicillium and Aspergillus species contributed more to ergosterol than Eurotium species. Ergosterol levels in sound canola were between 1.46 and 1.67 mg/g, whereas levels above 2 mg/g indicated significant levels of spoilage (Pronyk et al., 2006). Karaca and Nas (2006) examined ergosterol content of dried figs and found good correlation (r = 0.92) between aflatoxin and ergosterol in reject figs which were fluorescent, but no significant correlation with patulin content. Kadakal et al. (2005) found good correlation (r = 0.98) between ergosterol and patulin in apple juice and that both patulin (r = 0.99) and ergosterol (r = 0.99) were linearly related to the proportion of decayed apples used to make the juice. Ergosterol has been used to assess mould growth in cheese with variable results (Pecchini, 1997; Taniwaki et al., 2001a).

4.7 Estimation of Fungal Biomass

Ergosterol content has also been investigated as an indicator of the mycological status of tomato products. Battilani et al. (1996) found a significant correlation between ergosterol, Howard mould count (HMC) and fungal growth, but with a high level of uncertainty. Kadakal et al. (2004) found a linear relationship between degree of decay in tomato pulp and HMC (r = 0.97) and ergosterol (r = 0.96) and concluded that ergosterol has the potential to be used in quality assessment of tomatoes. Sio et al. (2000) described an improved method for extraction of ergosterol from tomato products. Other applications of ergosterol as a measurement of fungal biomass include estimation of mould spores in indoor air and aerosols (Miller and Young, 1997; Robine et al., 2005; Lau et al., 2006), estimation of wood decay by fungi (Eikenes et al., 2005) and estimation of fungi in soil and wetlands (e.g. Headley et al., 2002; Zhao et al., 2005). The ergosterol assay is reported to have a high sensitivity and, in contrast to the chitin assay, requires only 1 h for completion (Seitz et al., 1979). Despite its limitations, it appears to be a useful indicator of fungal invasion of foods and to hold promise as a routine technique for quality control purposes.

4.7.3 Impedimetry and Conductimetry Metabolites produced by growth of microorganisms in liquid media alter the medium’s impedance and conductance. The use of changes in these properties as a measure of bacterial growth was suggested by Hadley and Senyk (1975) and was first applied to yeasts by Evans (1982) and to moulds by Jarvis et al. (1983). Most of the subsequent work on fungi has been carried out with yeasts, but the methodology is often applicable to moulds also. During the 1980s there were a number of studies aimed at optimising media for inducing detectable and reproducible changes in either conductance or capacitance during fungal growth (see Pitt and Hocking, 1997). Watson-Craik et al. (1989, 1990) studied 27 mould species on a wide range of both commercially available and specially ppared media and concluded that conductance and capacitance were both medium and species specific. The use of media high in ammonium ions and glucose, with added yeast extract and peptone, decreased the

37

influence of product variability and induced higher conductance changes (Owens et al., 1992). An impedimetric method for detection of heat resistant fungi in fruit juices was described by Nielsen (1992). The detection limit in artificially contaminated juices was one Neosartorya ascospore per millilitre, detectable in 100 h. Huang et al. (2003) reported an impedance method for detection of bacteria and fungi in bottled water which shortened the detection time from 5 days to 27.1 h for fungi and from 48 to 11.3 h for bacteria. Although impedimetry and conductimetry promised to be effective rapid methods when used under well-defined conditions for a specific purpose with a particular kind of food, the methodology does not appear to have been broadly taken up for food mycology applications.

4.7.4 Adenosine Triphosphate (ATP) ATP has also been suggested as a measure of microbial biomass because bioluminescence techniques provide a very sensitive assay (Jarvis et al., 1983). Provided that background levels of ATP in plant or other cells are very low, or that microorganisms can be effectively separated from such other materials, the method has some potential as a microbial assay. A good correlation was shown between ATP production and viable counts of six species of psychrotrophic yeasts grown in pure culture (Patel and Williams, 1985). The effective detection of low levels of yeasts in carbonated beverages by ATP has also been reported (LaRocco et al., 1985). However, living plant cells contain high levels of ATP and fungi are often very difficult to separate from food materials. Moreover, extraction of molecules from fungal cells is notoriously difficult, so this potential may be difficult to realise in food mycology. The most widespad application of ATP bioluminescence in the food industry is for monitoring hygiene of surfaces in food production facilities (Easter, 2007).

4.7.5 Fungal Volatiles Methods for detection and characterisation of fungal volatiles are finding increasing applications in

38

4 Methods for Isolation, Enumeration and Identification

of volatile compound production as an indicator of mould deterioration in grains has been extensively assessed and reviewed (Kaminski and Wasowicz, 1991; Schnu¨rer et al., 1999; Magan and Evans, 2000; Paolesse et al., 2006; Balasubramanian et al., 2007). Fungal volatiles can be used to detect potential mycotoxin contamination, to discriminate between fungal species (Sunesson et al., 1995; Keshri et al., 1998) and even between toxigenic and non toxigenic strains of particular fungi (Sahgal et al., 2007). Karlshøj et al. (2007a) used an electronic nose to differentiate between closely related Penicillium species (P. camemberti, P. nordicum, P. paneum, P. carneum, P. roqueforti and P. expansum) from cheese. Volatile profiles can be used to pdict mould spoilage in bakery products (Vinaixa et al., 2004; Marı´ n et al., 2007a) and to detect and differentiate between toxigenic and non toxigenic P. verrucosum strains in bakery products (Needham and Magan, 2003). Volatile profiles have also been used to differentiate between toxigenic and non toxigenic Fusarium strains (Keshri and Magan, 2000; Demyttenaere et al., 2004), to identify mycotoxins (aflatoxins, ochratoxin A and deoxynivalenol) in durum wheat (Tognon et al., 2005) and to detect and quantify ochratoxin A and deoxynivalenol in barley (Olsson et al., 2002). This technology has also been applied to pdict the psence of P. expansum and patulin in apple products (Karlshøj et al., 2007b) and to detect and discriminate diseases of potato tubers (Kushalappa et al., 2002) and stemend rot and anthracnose in mangoes (Moalemiyan et al., 2006). Electronic nose technology has also been used for early detection of moulds in libraries and archives (Pinzari et al., 2004).

4.7 Estimation of Fungal Biomass

Enzyme-linked immunosorbent assay (ELISA). The pparation of antigens from three common foodborne fungi (Penicillium aurantiogriseum, Mucor racemosus and Fusarium oxysporum) was described by Notermans and Heuvelman (1985). Preparation of immunoglobulin antibodies against these antigens was followed by development of an ELISA assay. Fungi were detected in both unheated and heat processed foods by this method. Antigens were relatively genus specific: the M. racemosus antigens reacted with other Mucor and Rhizopus species, and the Penicillium antigen reacted with the Aspergillus species tested. It was subsequently shown that the Penicillium antigen reacted with 43 of 45 Penicillium species tested and that antigen production correlated with mycelial weight and was unaffected by culture conditions, medium, temperature and aw (Notermans et al., 1986). The Penicillium antigen also reacted with Aspergillus flavus and the level of antigen correlated with aflatoxin production (Notermans et al., 1986). ELISA techniques have also been studied as a potential replacement for the Howard mould count. Antigens from tomato moulds (Alternaria alternata, Geotrichum candidum and Rhizopus stolonifer) were used to produce an ELISA test sensitive to 1 mg/g of mould in tomato. A correlation was observed between antigen formation and mould added to tomato puree, while background interference was very low (Lin et al., 1986). The method was tested against a broader range of foods, with encouraging results (Lin and Cousin, 1987). Robertson and Patel (1989) improved the sensitivity of the method for tomato paste by using a polyclonal antibody against Botrytis cinerea, Mucor piriformis and Fusarium solani in addition to the three species used by Lin et al. (1986). ELISA based methodology has been reported for the detection of Fusarium species in corn (Meirelles et al., 2006), cornmeal (Iyer and Cousin, 2003) and grain (Rohde and Rabenstein, 2005). Correlations with other measures of fungal growth (ergosterol, colony count, mycotoxin levels) were variable. Aspergillus species have also been examined as targets for immunological detection because of their importance in mycotoxin contamination. ELISA detection of A. ochraceus in wheat (Lu et al., 1995), coffee powder, chilli powders and poultry feed (Anand and Rati, 2006) has been investigated, with reasonable correction with other parameters

39

40

Schwabe et al. (1992) compared the latex agglutination assay with ergosterol production for detection of Penicillium, Aspergillus and Fusarium species in pure culture. They concluded that the two methods were comparable for Penicillium and Aspergillus but that ergosterol was more sensitive for Fusarium. In food samples, both the latex agglutination test and ergosterol were effective means of detecting mould growth, but no clear correlation existed in values obtained by the two methods. Kesari et al. (2004) used a latex agglutination test to detect teliospores of Karnal bunt (Tilletia indica) in single grains of wheat and were able to detect a few as 750 teliospores, which they reported as suitable for single seed analysis. Fluorescent antibody techniques. Fluorescent antibody techniques have also been used directly for the detection of mould in foods. Warnock (1971) detected Penicillium aurantiogriseum in barley by this method, while Robertson et al. (1988) used antisera from five fungi to visualise moulds and simplify their detection in the Howard mould count technique.

4 Methods for Isolation, Enumeration and Identification

4.8 Identification Media and Methods

Sequencing of the ITS region along with ”housekeeping genes” such as calmodulin, b-tubulin and elongation factor 1-a is now commonly applied for purposes of identification and phylogenetic analysis of important food spoilage and mycotoxigenic fungi in the genera Aspergillus (Varga, 2006; Geiser et al., 2007; Peterson, 2008), Penicillium (Peterson, 2004, 2006; Samson et al., 2004b; Wang and Zhuang, 2007; Serra et al., 2008) and Fusarium (O’Donnell et al., 2004; Scott and Chakraborty, 2006; Leslie et al., 2007). Despite the apparent power of molecular techniques, they need to be applied with some caution, particularly when comparing DNA sequences with those in the publicly available databases to identify yeast or mould isolates. Correct identification relies on the database sequences having the correct name attached to them by the depositor, which is not always the case. If the identification makes sense, the percent homology is 98% or greater and the number of base pairs on which the homology is scored is high, then the answer is probably correct.

4.8 Identification Media and Methods 4.8.1 Standard Methodology The identification keys in this book are based primarily on the standardised procedure described for the identification of Penicillium species by Pitt (1979b). Cultures are grown for 7 days on three standard media at 258C, and on one of these at 5 and 378C also. The three Media are Czapek yeast extract agar (CYA; Pitt, 1973) used at all three temperatures; malt extract agar (MEA; Raper and Thom, 1949) and 25% glycerol nitrate agar (G25N; Pitt, 1973). Their formulae are given in the Media Appendix. Preparation time of CYA and G25N is reduced by the use of Czapek concentrate (Pitt, 1973), which is added to the media at the rate of 1% of the aqueous portion. As media ingredients have become more purified in recent years, difficulties with extent and colour of sporulation on CYA have been encountered, especially with some Penicillium species. To overcome this problem, Czapek concentrate has been reformulated (Pitt, 2000) by the inclusion of traces of zinc and copper (Smith, 1949) (see Media Appendix).

41

4.8.3 Inoculation As shown in Fig. 4.3, Petri dishes of CYA and MEA for incubation at 258C are inoculated with a single culture at three points, equidistant from the centre and the edge of the plate and from each other. Plates of the other media are inoculated with two points per culture, as illustrated. With some fungi, especially Penicillium and Aspergillus, it is important to minimise colonies from stray spores. The most satisfactory technique is to inoculate plates with spores suspended in semisolid agar (Pitt, 1979b). Dispense 0.2-0.4 ml of melted agar (0.2%) and detergent (0.05%), such as polysorbitan 80 (Tween 80), in small vials and sterilise. To use, add a needle point of spores and mycelium to a vial and mix slightly. Then, before flaming the needle, use it to stab inoculate the 58C plate;

42

4 Methods for Isolation, Enumeration and Identification

Fig. 4.3 Schematic of regimen used for culturing fungal isolates for identification

residual spores on the needle make a good inoculum. Next, take a sterile loop, mix the vial contents thoroughly and inoculate the standard plates. Used vials can be sterilised by steaming and reused several times before being washed or discarded.

4.8.4 Additional Media and Methods The plating regimen outlined above can be used to identify most of the fungi described in subsequent chapters of this book. Some exceptions exist, because certain genera either grow poorly or fail to sporulate on the standard media. As noted earlier in this chapter, fastidious xerophiles are identified on MY50G agar. Eurotium species, traditionally identified on Czapek agar with 20% sucrose, are identified here on Czapek yeast extract agar with 20% sucrose (CY20S; see Appendix). Trichoderma species are best identified on potato dextrose agar (PDA) after a relatively short incubation time (3-4 days), as the structures tend to autolyse as cultures mature. Penicillium subgenus Penicillium. Many species classified in subgenus Penicillium are morphologically similar, and identification using traditional morphological techniques remains difficult. These

Penicillia are very common in foods, and many produce mycotoxins, so correct identification is often critical. Species within subgenus Penicillium fall into two groups: those with an affinity for proteinaceous foods and those which grow more vigorously in foods high in carbohydrate. Frisvad (1981, 1985) introduced creatine sucrose agar (CREA) to permit differentiation between these two groups. Creatine (as the sole nitrogen source) permitted growth of the former group while inhibiting the latter. Incorporation of bromocresol purple enabled visualisation of pH changes, either acid or alkaline, depending on the creatine or sucrose metabolism of a particular species. However, discrimination between positive and negative responses was not always clear, and the main tabulation of species reactions to CREA (Table 1 in Frisvad, 1985) was difficult to interpt. Frisvad (1993) subsequently produced a number of variations of CREA, including acid and neutral pH formulations and substitution of sucrose with fructose or lactose. Although the merits of each formulation were discussed, no firm recommendation resulted from this exercise. Pitt (1993a) modified Frisvad’s strongly alkaline CREA medium by studying several sucrose and creatine concentrations over a wide pH range. The result was neutral creatine sucrose agar (CSN), a medium producing eight different reactions among

4.8 Identification Media and Methods

the 20 Penicillium subgenus Penicillium species tested. When included in the normal plating regimen for identification of Penicillium cultures from the subgenus Penicillium, CSN provides a very useful aid to distinguishing between these difficult and closely related species. See Chapter 7 for details of use of CSN, its reactions and interptation. Its formulation is given in the Media Appendix. Dematiaceous Hyphomycetes. The natural habitats for many dematiaceous Hyphomycetes are plants or plant material, so finding suitable laboratory media and conditions to induce typical sporulation can be difficult. Alternaria, Curvularia, Stemphylium and Ulocladium species are best identified from dichloran chloramphenicol malt extract agar (DCMA; Andrews, 1992b) plates incubated for 7 days at 258C under lights. Conidial characteristics of Bipolaris species vary with type of medium. Species described here are best identified from tap water agar (TWA) containing a natural substrate such as sterilised wheat or millet seeds, or wheat straw. Drechslera species will grow well on DCMA but sporulation is poor. The best sporulation is achieved with V-8 juice (V-8 J) agar or TWA with one of the natural substrates mentioned above, incubated at 258C for 7-10 days under lights with a 12 h photoperiod. Light/dark periodicity is important as Drechslera cultures require light to produce conidiophores and darkness to produce conidia, possibly due to light inactivation of flavine necessary for conidial formation (Knan, 1971; Platt et al., 1977). For the identification of Trichoderma and Fusarium species, potato dextrose agar (PDA) is used. Fusarium species also require additional methods and media as outlined below.

4.8.5 Identification of Fusarium Species Fusarium isolates exhibit unusually high variability in colony morphology and also may deteriorate rapidly in culture. Thus, they should be identified as soon as possible after initial isolation with a minimum of subculturing to avoid deterioration. It is common practice to ppare cultures of Fusaria from single spores for growth on identification media, as this reduces both of these problems.

43

Single sporing. The technique for pparing single spore cultures is as follows (Nelson et al., 1983; Leslie and Summerell, 2006). Pour about 10 ml of 2% water agar into unscratched glass or plastic Petri dishes and allow to dry, either by holding the plates at room temperature for several days or by placing them inverted in an oven at 37-458C for about 30 min. Prepare a suspension of conidia in a 10 ml sterile water blank so that it contains 1-10 spores per low-power (10) microscope field when a drop from a 3 mm loop is examined on a slide. With experience, this concentration can be gauged simply by observing the turbidity of the suspension. Pour the suspension of spores onto a dried water agar plate, drain off the excess liquid and incubate in an inclined position at 20-258C for 18-20 h. After incubation, open the Petri dish, shake off any accumulated moisture droplets and examine under a stereomicroscope using transmitted light. The germinating conidia should be visible under 25 magnification. A dissecting needle with a flattened end and sharpened edges is used to cut out small squares of agar containing single, germinating conidia. These single conidia are then transferred on the agar blocks to the desired growth medium. If the original culture is contaminated with bacteria, a drop of 25% lactic acid may be added to the water blank. Allow this acidified spore suspension to stand for 10 min before pouring onto a water agar plate. Germination of acid-treated Fusarium conidia may be delayed by 24 h or more. Media. Two media have been used in this book for the identification of Fusarium isolates: potato dextrose agar (PDA) for colony characteristics and colour and dichloran chloramphenicol peptone agar (DCPA) for the development of diagnostic macro-, micro- and chlamydoconidia. A third medium, carnation leaf agar (CLA), is recommended by some Fusarium specialists for Fusarium cultivation and identification (Nelson et al., 1983; Leslie and Summerell, 2006). CLA is an excellent medium, on which most Fusarium species readily produce their diagnostic macroconidia. Production of macroconidia on DCPA is usually comparable with that on CLA, but microconidia and chlamydoconidia are often more plentiful on CLA due to greater production of aerial hyphae. DCPA is used in the psent work rather than CLA,

44

however, because dried, gamma-irradiated carnation leaves are difficult to obtain in many localities. Inoculation and incubation. For identification by the methods used in this book, single spore cultures of Fusarium isolates should be ppared on agar blocks as outlined above, inoculated, one per plate, onto two plates each of PDA and DCPA and incubated at 258C for 7 days. Inpidual plates may be used for each medium, or alternatively pided plates may be used, with one medium on each half of the plate. Illumination during incubation is essential for the production of macroconidia. The light source may be diffuse daylight (not direct sunlight) or light from a bank of fluorescent tubes. A photoperiod of 12 h per day is normally used. Alternating temperatures of 20 and 258C have been recommended (Nelson et al., 1983; Leslie and Summerell, 2006) but are not essential. A simple light bank may be constructed from a standard 40 watt fluorescent fixture with two cool white tubes, suspended 0.5-l m above the laboratory bench or shelf supporting the cultures. The addition of a black light tube (e.g. Philips TL 40 W/80 RS F40BLB) is also desirable and in some cases essential to induce macroconidial or chlamydoconidial production.

4.8.6 Yeasts Yeast identification systems. Identification of foodborne yeasts remains a difficult task, as colony characteristics and microscopic morphology are of limited value. Generally it has been necessary to use biochemical and physiological tests such as fermentation of carbohydrates, assimilation patterns for a range of carbon and nitrogen sources and growth at various temperatures. Details of these methods and media may be found in Kurtzman and Fell (1998), Kurtzman et al. (2003) or Barnett et al. (2000). Molecular methods (see below) are being used increasingly in yeast identification, replacing time consuming biochemical testing. Identification using systems based on biochemical and physiological testing is complex and time consuming. However, a number of attempts have been made to assist those who wish to persevere with yeast identification. Several simplified systems

4 Methods for Isolation, Enumeration and Identification

have been published in the literature, and both automated and manual yeast identification systems are now commercially available. Dea´k and Beuchat (1987) published a simplified identification key (SIK) which included 215 species of foodborne yeasts. They subsequently modified their system, restricting it to the 76 species most frequently occurring in foods (SIM), and reported that it was much more successful than the API 20C system (BioMe´rieux, Marcy-l’Etoile, France) for identification (Dea´k, 1992). The SIM uses only two Petri dishes and three test tubes to examine each strain for ability to assimilate 10 carbon sources, fermentation of glucose, assimilation of nitrate and splitting of urea. These biochemical tests are supplemented by morphological observations. SIM separates the yeasts into six groups by a dichotomous key utilising the results of five key tests. Further tests are used to differentiate the yeasts in these six groups using secondary dichotomous keys. A dichotomous key to 25 common species of foodborne yeasts was published by Smith and Yarrow (1995), who used 17 biochemical and physiological tests to distinguish the species. Of the commercial systems available, the most widely used for foodborne yeasts are the Biolog, which is an automated system, and the BioMe´rieux ID32C yeast identification strips which can be read manually or automatically using the ATB system. BioMe´rieux also markets the fully automated VITEK 2 system, based on a card containing 64 tests. However, the database comprises only 46 clinically important species (Aubertine et al., 2006) and so has limited application to foodborne yeasts. The Biolog (Biolog Inc., Hayward, CA, USA; http://www.biolog.com/microID.html) utilises a yeast identification test panel (YT MicroPlateTM) consisting of a matrix of 8 12 wells. The first three rows contain 35 carbon source oxidation tests using tetrazolium violet as an indicator of oxidation. The next five rows contain carbon assimilation tests which are scored turbidimetrically against a negative control panel containing only water. The last row contains two carbon sources and tests for the co-utilisation of various carbon sources with D-xylose. The hardware (Biolog MicroStation Reader) consists of an automated plate reader coupled with a computer, which interpts the

4.9 Examination of Cultures

results and compares them with the resident database which currently includes 267 species. Manual interptation of the Biolog plates is not recommended. This system has been designed with the food industry in mind, and the database contains all the common foodborne yeasts, unlike other systems which are usually aimed at the clinical market. The ATB ID32C system (BioMe´rieux, Marcyl’Etoile, France) is an automated system utilising the BioMe´rieux ID32C yeast identification strips. These strips contain 30 assimilation tests plus a positive (glucose) and a negative control well, all of which are inoculated with a yeast suspension of specified density. The strips are incubated at 308C for 48-72 h. As with the Biolog, the ATB automated system consists of a plate reader attached to a computer. The database associated with the ATB system contains 63 yeast species. The BioMe´rieux ID32C strips can be read manually, and the results enable identification of yeasts using published keys or computer identification programmes such as that of Barnett et al. (1996). This system may be used with reasonable success, particularly when the test results from the ID32C strips are supplemented with extra tests (glucose fermentation, urease production, nitrate utilisation, growth in 0.5 and 1% acetic acid and in 50 and 60% glucose, growth at 378C, production of pseudohyphae, ascospore formation and morphological observations) to give a more comphensive base for the identification. Growth in 0.5 and 1% acetic acid indicates pservative resistance, and growth in 50 and 60% glucose gives an indication of ability to grow at reduced aw. Both these parameters are important in determining a yeast’s ability to cause spoilage in particular products. Identification using DNA sequencing is increasingly becoming the method of choice, as extensive databases such as GenBank are freely available for identification purposes. The 600- 650 nucleotide D1/D2 region of the large subunit (26S) ribosomal DNA is the most widely targeted section of the genome, and sometimes the ITS region may also be used (Kurtzman et al., 2003). Sequencing of the D1/D2 region, along with some supplementary physiological and biochemical tests, is the identification method currently used in our laboratory.

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Even using the available systems, identification of yeasts still requires some specialist knowledge and interptation and remains time consuming. Our experience indicates that no more than 12 species of spoilage yeasts are of real concern in foods. It is possible to differentiate these few species by relatively simple techniques, i.e. colony and microscopic morphology, growth on the standard media used for filamentous fungi, growth on other media which test for pservative resistance, ability to use nitrate as a nitrogen source and adaptation to high NaCl concentrations. Details of these techniques are given in Chapter 10.

4.9 Examination of Cultures As noted above, all cultures for identification should first be grown on the standard regimen described earlier. After 7 days incubation, the following examination should be carried out and then the general key to fungi in Chapter 5 should be used. That key will be of assistance even in the event of cultures failing to grow under one or more of the standard conditions.

4.9.1 Colony Diameters Measure the diameters of macroscopic colonies in millimetres from the reverse side (Fig. 4.4). Microscopic growth or germination at 58C is assessed by low power microscopy (60-100), by putting the 58C Petri dish on the microscope stage and examining by bright field transmitted light. Growth at 378C is assessed macroscopically only; germination of spores at 378C is an unreliable character.

4.9.2 Colony Characters Colony appearance can be judged by eye or with a hand lens, but examination is more effective if a stereomicroscope is used. Magnifications in the range of 5-25 are the most useful. Characters such as type and location of sporing structures and

46

4 Methods for Isolation, Enumeration and Identification

Fig. 4.4 Technique for measuring colony diameters by transmitted light

extent of sporulation are best gauged with the stereomicroscope. Reflected light is usually more effective than transmitted light. To determine colony colours, examine colonies by daylight or by daylight-type fluorescent light. In some genera, reference to a colour dictionary is helpful. The Methuen ”Handbook of Colour” (Kornerup and Wanscher, 1978) has been used in this work and is highly recommended.

4.9.3 Preparation of Wet Mounts for Microscopy Fungi should always be examined microscopically as wet mounts rather than fixed and stained like bacteria. To ppare a wet mount, use an inoculating needle to cut out a small portion of the colony which includes sporing structures. Examination with the stereomicroscope can be an invaluable aid here. For freely sporing fungi with little mycelium, cut a piece of colony near the edge, where fruiting structures are young and spore numbers not excessive. Take structures which may enclose spores, i.e. cleistothecia, from near colony centres, where the probability of mature spores is highest. If the only differentiated parts of the colony appear to be

buried in the agar, e.g. pycnidia, take a sample of these with a small piece of the agar. Float the cut colony sample from the needle onto a slide with the aid of a drop of 70% ethanol. It may be necessary to tease out the specimen with the needle and the corner of a cover slip (square cover slips are best). Fungal specimens may be highly hydrophobic: the ethanol helps to wet the pparation, minimising the amount of entrapped air. When most of the ethanol has evaporated, add a drop of lactic acid (for phase or interference contrast optics) or lactofuchsin stain (see below) for bright field. Add a cover slip; if necessary remove excess liquid from the pparation by gently blotting with facial tissue or similar absorbent paper. The pparation is now ready for examination.

4.9.4 Staining A wide variety of stains are in use for mycological work. However, most are time consuming to ppare, or to use, or are slow to act, because fungal walls and spores are highly resistant to stains. By far the most effective stain for use in food mycology is lactofuchsin (Carmichael, 1955), which suffers from none of these faults. It consists of 0.1% acid fuchsin

4.9 Examination of Cultures

dissolved in lactic acid of 85% or higher purity. Young actively growing fungal structures are pferentially coloured bright pink, so sporing structures can usually be readily distinguished against a background of older mycelium. Cleistothecial initials, developing asci and maturing ascospores are also seen more readily in pparations stained with lactofuchsin. Like most other mycological stains, lactofuchsin is corrosive. Take care to clean it off microscope parts or skin! Be especially careful of the objective faces, because lactic acid will slowly corrode the relatively soft glass used in lenses.

47

equipped with an eyepiece micrometer, which is essential for measuring dimensions of spores and sporogenous structures. In the examination of fungal mounts, it is stressed that it is most important to use low power optics before succumbing to the temptation to use oil immersion. The principal reason is that fungal pparations usually remain as small clumps and do not disperse as bacteria do. Only under low power is the search for the optimal area of the slide for the observation of fruiting structures likely to be rewarded. Once a suitable area is located under the 10 or 16 objective, move to the 40. This should be the lens most used; microscope optics are such that only the finest details of ornamentation can be observed more effectively under oil immersion than at this magnification. Aligning the microscope. Correct alignment of the microscope is essential, so that its resolution is as high as possible and it can be used for long periods without discomfort. An incorrectly aligned microscope will lead to poor observation, discomfort, fatigue, headache and eye strain. A person of normal visual acuity should be able to use a correctly aligned instrument throughout a whole working day without discomfort. The steps to correctly align a microscope are given below. They should be read in conjunction with the microscope manufacturer’s instructions. 1. Mount a slide on the stage and bring it into approximate focus. If a ppared slide is not available, a slide marked with a marking pen or ink line is a satisfactory substitute. 2. Close the microscope’s field diaphragm (the one at the microscope base nearer the light source). The image of the diaphragm opening should now be visible in the microscope field. If it is, first focus it with the condenser focusing knob and then centre it in the field with the condenser centring screws. If the diaphragm opening cannot be seen, first rack the condenser up and down and watch to see if the opening becomes visible; if it does not, rack the condenser to its highest position and then slowly open the field diaphragm until the opening comes into view. Centre the diaphragm approximately and proceed as above. 3. For bright field optics, the condenser diaphragm should be adjusted each time the objective power is changed. Remove one ocular; close the

48

condenser diaphragm so that the field seen down the open tube is about two-thirds its maximum size. With phase contrast and interference contrast systems, this adjustment is less critical. The pceding steps align the microscope itself and should be checked frequently. If optimal illumination is desired, each step should be carried out for each new slide and each objective change. As a routine habit, the whole process should take only a few seconds. If the available microscope is not equipped with a built-in light source, a field diaphragm and a fully centring condenser, it is unlikely to be satisfactory for the identification of small spored fungi such as Penicillium species. The following steps are designed to align the observer with the microscope, compensating for inpidual differences in sight. Provided settings on the microscope are remembered, these steps need be carried out only occasionally, to check that visual acuity has not altered. Different settings will be needed for an inpidual with and without spectacles or contact lenses. 4. Assuming the microscope is binocular, pull the oculars out to their greatest distance apart and then, while watching a focused field, move them gradually together until a single circular field is seen without strain or head movement. Note the distance on the scale between the oculars; this is the inpidual’s interpupillary distance. Repeat this operation two or three times until satisfied that the correct distance has been found. This distance should always remain the same and be similar on any microscope. 5. Under the 40 or 100 objective, locate a tiny, readily recognised point on the slide and focus on it. Take a piece of white card and place it between the focusing ocular and the corresponding eye. Leave the eye open. Now focus the tiny point with the other eye, carefully, with the microscope fine focus. Next, transfer the white card to the other ocular and, using the focusing collar beneath the ocular, refocus the tiny point. Remove the card and note the setting on the scale. Repeat until satisfied the correct setting has been found. 6. On some microscopes, the eyepiece micrometer can be focused independently. Use the focusing system on the ocular itself to focus the micrometer. Note the setting on the scale on the side of the ocular.

4 Methods for Isolation, Enumeration and Identification

Always check the settings on the microscope before use and after making measurements with the micrometer. It is very easy to upset the ocular alignment when measuring.

4.10.1 Lyophilisation For unstable cultures, and indeed for the long term storage of any food spoilage fungi, freeze drying or lyophilisation is probably the best method of pservation. Many commercial systems are now available for carrying out this process. A satisfactory menstruum for lyophilisation of most fungi is l.5 normal strength reconstituted nonfat milk powder (15% in distilled water). For fungi with hydrophobic conidia, such as Aspergillus and Penicillium, a small amount of detergent (0.05%) such as polysorbitan 80 (Tween 80) should be added to the milk. Dispense the milk in 10 ml lots in small tubes or 12.5 ml (0.5 oz) McCartney bottles, and sterilise by steaming for 20 min on three successive days (the Tyndallisation process). The milk must be stored at 208C or above between steamings, to permit bacterial spores to germinate.

4.10 Preservation of Fungi

4.10.2 Other Storage Techniques A variety of systems other than lyophilisation have been proposed for long term storage of fungi. Of these, liquid nitrogen storage has found most acceptance with major culture collections. This type of storage appears to be superior to any other for plant pathogens and fungi which will not sporulate in pure culture. However, liquid nitrogen systems are expensive to establish and maintain and are only suitable for large collections. Freezer units which run at very low temperatures (-808C or below) are available and are well suited to the needs of culture collections. In our laboratory we routinely store cultures at -808C, using glycerol (60-80%) as a cryoprotectant. Spore suspensions are ppared by taking conidia or ascospores from a freely sporulating sector of the colony, dispersing them in the glycerol then freezing in screw capped

49

cryovials. An 80% solution of glycerol remains viscous at -808C, which enables cultures to be removed from the freezer for subculture without the need for defrosting. For smaller laboratories that do not have access to lyophilisation, liquid nitrogen or ultra-low temperature storage, some simple techniques exist that can be used to maintain fungal cultures over relatively long periods (e.g. 1-10 years) without the need for subculturing. Water storage. A simple and inexpensive method of fungal culture pservation is storage of agar blocks in water (Smith and Onions, 1994). Small agar blocks (7-10 mm2) are cut from the growing margin of a young fungal colony and placed in sterile water in a bottle such as a Bijou bottle (6.25 ml or 0.25 oz McCartney bottle). The rubber lined cap is screwed down and the bottles stored in a cool room (1-108C). Cultures may be revived by removal of a block and placing it on a suitable growth medium. Using this method, cultures have been reported to remain viable and retain their characteristics for up to 7 years (Boeswinkel, 1976; Smith and Onions, 1994). Some yeasts may be maintained by storing as suspensions in water (Kirsop, 1984). Growth from a late logarithmic slant culture is suspended in sterile distilled water and transferred to a sterile container so that 90% of the volume is filled with the suspension. Containers are stored at room temperature. Survival of some Candida, Saccharomyces, Cryptococcus, Rhodotorula and Schizosaccharomyces species for up to 4 years has been reported (Kirsop, 1984). Silica gel storage. Many fungal cultures may be maintained for long periods (often more than 10 years) by drying spore suspensions onto silica gel (Smith and Onions, 1983; Smith, 1984). This method is not suitable for mycelial cultures, but can be used with some success for yeasts (Kirsop, 1984). As silica gel liberates heat when moistened, the technique depends on keeping the cultures cool enough to avoid damaging the spores during pparation. Medium grain plain (non-indicating) silica gel of 6-22 mesh is placed in suitable glass bottles (Bijou or McCartney bottles) to a depth of about 1 cm and sterilised, either by dry heat at 1808C for 2-3 h or by autoclaving at 1218C for 15 min. Autoclaved silica gel must be thoroughly

50

dried in an oven before use. Bottles are pcooled by placing in a tray of ice or refrigerating for 24 h, then transferring to an ice tray for inoculation. Suspensions of fungal spores or yeasts are ppared in sterile skimmed milk (as for lyophilisation, above) and the suspension added to the cooled silica gel to wet three-quarters of it. The bottles and gel are allowed to dry at room temperature for 10-14 days with caps slightly loosened. Caps are then screwed down and the bottles stored at 48C (storage at room temperature is also satisfactory) in air-tight containers over indicating silica gel to absorb any moisture. Cultures are revived by shaking a few crystals of silica gel onto a suitable growth medium (broth culture may be better for yeasts). Survival varies according to species or even strain, but survival of yeasts for up to 5 years (Kirsop, 1984) and fungi for more than 10 years (Smith and Onions, 1983) has been reported.

4.11 Housekeeping in the Mycological Laboratory Like any other microbiological laboratory, a mycological laboratory should be kept in a clean condition. Discard unwanted cultures regularly and dispose of them by steaming or autoclaving. Wipe bench tops regularly with ethanol (70-95%). Floors should be wet mopped or polished only with machines equipped with efficient vacuum cleaners and dust filters. Where possible store food and plant materials away from the laboratory. Open Petri dishes carefully. Use small inocula on wet needles. Transport Petri dishes to the stereomicroscope stage before removing lids. Do not bump cultures during transport. Contrary to popular belief, a well-run mycological laboratory is not a source of contamination to bacteriological laboratories. The air in a mycological laboratory should not carry a significant population of fungal spores. The reverse problem can occur, however, because bacteria multiply more rapidly than do fungi. Bacterial spores are often psent in food laboratories, readily infect fungal plates and can rapidly outgrow and inhibit fungal mycelia, especially at 378C.

4 Methods for Isolation, Enumeration and Identification

If for any reason fungal spore concentrations do build up in a laboratory and cause an unacceptable level of contamination, the air should be purified. The simplest technique is to spray the air throughout the laboratory with an aerosol before it is closed in the evening. Any aerosol spray, such as a room deodoriser or air freshener, is effective. Aerosol droplets entrain fungal spores very efficiently and carry them to the floor. A more drastic and effective treatment in cases of severe contamination is to spray a solution of 2% thymol in ethanol around the room and close it for a weekend. The spray is rather pungent, and while not harmful to humans, it effectively kills fungal spores and mites (see below). Do not leave cultures on benches before fumigation.

4.11.1 Culture Mites A major hazard in growing and maintaining fungal cultures is the culture mite. Many species of mites live on fungal hyphae as their main or sole diet in nature and find culture collections an idyllic environment. Mites crawl from culture to culture, contaminating them with fungi and bacteria as they go or, given long enough, may eat them out entirely. Mites are very small (0.05-0.15 mm long), usually just visible to the observant naked eye. They are arachnoids, related to spiders, and hermaphroditic. Each mite leaves a trail of eggs about half adult size as it goes. Eggs hatch within 24 h and reach adulthood within 2 or 3 days. The damage an unchecked mite plague can do to a fungal culture collection has to be experienced to be believed. The most common sources of mites are plant material, soil, contaminated fungal cultures and mouldy foodstuffs. Mites can also be carried on large dust particles. Building work near a laboratory almost always induces a mite infestation. The avoidance of losses due to mites requires constant vigilance. Always watch for telltale signs, such as contaminants growing around the edges of a Petri dish, a ”moth-eaten” appearance to colonies or ”tracks” of bacterial colonies across agar. Examination of suspect material or cultures under the stereomicroscope will readily reveal the psence of mites and mite eggs.

4.11 Housekeeping in the Mycological Laboratory

Adult mites are rapidly killed by freezing, and mite eggs will only survive 48-72 h at -208C. Cultures contaminated by mites can often be recovered by freezing for 48 h, then subculturing from uninfected portions of the culture with the aid of the stereomicroscope. Suspect food and other samples being brought into the laboratory should also be frozen for 24 h to destroy mites before enumeration or subculturing is carried out. Infestation by mites can be minimised by good housekeeping, i.e. by avoiding accumulation of dust or old cultures in the laboratory. It is also good practice to handle and store food and plant samples well away from areas where fungi are inoculated and incubated. To control a mite infestation, remove all contaminated material, including cultures. Freeze Petri dishes and culture tubes which must be recovered; autoclave, steam or add alcohol to all others. Clean benches thoroughly with sodium hypochlorite (household bleach) or 70% ethanol. Incubators can be disinfested with aerosol insecticides.

4.11.2 Problem Fungi There are three fungal invaders which should be watched for carefully in a food mycology laboratory: Aspergillus fumigatus, Rhizopus stolonifer and Chrysonilia sitophila. The first is a human pathogen; the others can cause a contamination chain which is difficult to break. Aspergillus fumigatus may cause invasive aspergillosis in the lungs or serious allergenic responses in some inpiduals. It is sound practice to immediately kill and discard cultures of this fungus as soon as it is recognised. On no account should it be used for experimental studies in food spoilage or biodeterioration without pcautions to pvent dissemination of spores. The morphology of A. fumigatus is described in detail in Chapter 8, but it is readily recognisable in the unopened Petri dish:

colonies are low, dull blue and broadly spading, with a velvety texture;

growth is very rapid at 378C, covering a Petri dish in 2 days;

long columns of blue conidia are readily seen under the stereomicroscope.

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4.11.3 Pathogens and Laboratory Safety While it must be said that any fungus which is capable of growth at 378C is a potential mammalian pathogen, the physiology of the healthy human is highly resistant to nearly all of the fungi encountered in the food laboratory. Nevertheless, fungi which can grow at 378C should be treated with caution. In particular, the habit of sniffing cultures is to be avoided wherever possible. It is true that odours produced by fungi have been used quite frequently as taxonomic criteria, especially in older

52

publications, but their subjective and ephemeral nature makes them of little value for this purpose, and the risks involved are serious. Some laboratories regard fungal volatiles as such a serious risk that cultures such as Penicillium and Aspergillus species are handled in a biohazard cabinet. Many types of fungal spores are allergenic or carry mycotoxins. Inhalation of fungal spores should be avoided as far as possible. Of the fungi described in this book, only the Aspergilli normally pose any direct threat to health. Aspergillus fumigatus has already been mentioned, and care should also be taken when handling cultures of Neosartorya species, which are closely related to

4 Methods for Isolation, Enumeration and Identification

A. fumigatus and also grow prolifically at 378C. Other Aspergillus species, particularly A. flavus, A. niger and A. terreus, have been isolated from human specimens from time to time. For more details, see de Hoog et al. (2000). These species appear to be mainly opportunists and pose little threat to healthy people. Careful handling and good housekeeping are all that are required. However, immunocompromised inpiduals are in a different category. It is increasingly evident that such people have little resistance to fungal infection. Persons suspected to be immunocompromised, regardless of the cause, should not work in a mycology laboratory nor indeed be permitted knowingly to enter one.

Chapter 5

Primary Keys and Miscellaneous Fungi

Principles underlying fungal classification have been outlined in Chapter 3, including a brief overview of the relevant pisions of the Kingdom Fungi and their principal methods of reproduction. Some further detailed information is necessary in this chapter to assist in the use of the keys which follow. Ascomycetes. As discussed in Chapter 3, Ascomycetes produce ascospores in asci (Fig. 3.2). One genus, Byssochlamys, produces asci which are unenclosed; all other genera of relevance here produce asci in some kind of fruiting body, or ascocarp. The two kinds of ascocarp commonly seen in food spoilage fungi, the cleistothecium and the gymnothecium, have been described and illustrated in Chapter 3 (Fig. 3.3). Both types of ascocarp are usually pale or brightly coloured, not dark, and release ascospores by rupturing irregularly. Of genera relevant here, cleistothecia are produced by Emericella, Eurotium, Eupenicillium, Monascus and Neosartorya, and gymnothecia by Talaromyces. A third class of ascocarp, less commonly encountered in foodborne fungi, is the perithecium. Perithecia have cellular walls like cleistothecia, but are distinguished by the psence of an apical pore or ostiole through which asci or ascospores are liberated; also asci are long and clavate with ascospores arranged linearly within them. In the one perithecial genus of interest here, Chaetomium, the perithecia are black and have stout hyphae attached to the walls (Fig. 5.10). Conidial fungi. The strictly conidial fungi, also known as Fungi imperfecti or Deuteromycetes or anamorphic fungi, possess an amazing variety of ways of producing conidia. Terminology for structures bearing conidia and for conidia themselves has

become astonishingly complex in recent years; fortunately most of it is not essential for the recognition of the genera discussed in this text. Terms which are important in the keys which follow are described below. A fundamental pision within the asexual fungi separates genera which form conidia aerially, grouped in the class Hyphomycetes, from those in which conidia are borne in some sort of enveloping body, the class Coelomycetes. Hyphomycetes. Fungi have developed seemingly endless ways of extruding or cutting off conidia, solitarily or in chains, from fertile cells which themselves may be borne solitarily or aggregated into more or less ordered structures. Hyphomycete taxonomy attempts to thread a way through this maze. In general, type and degree of aggregation of the fertile cells, and type of conidium, provides the basis for generic classification, while details of these characters and of spore size, shape and ornamentation are more commonly used to distinguish species. Features of conidia used in the keys in this work are length, septation, ornamentation and colour, particularly whether walls are light or dark. The method of conidium formation (ontogeny) is seldom emphasised here, because terminology is complex and distinctions may not be obvious. The principal point to note is the disposition of conidia: they may be borne solitarily, i.e. just one conidium per point of production; singly, i.e. successively from a single point, but unattached to each other; or in chains. Solitary conidia are borne on a relatively broad base and usually adhere to the fertile cell. Conidia formed in chains are usually extruded from a small cell of determinate length, often a

J.I. Pitt, A.D. Hocking, Fungi and Food Spoilage, DOI 10.1007/978-0-387-92207-2_5, Ó Springer ScienceþBusiness Media, LLC 2009

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phialide, which in most genera narrows to a distinct neck. Conidia borne singly may be extruded in this same manner, or be borne by extrusion from a pore in a hypha or fertile cell, or be cut off by hyphal fragmentation. Phialidic Hyphomycetes. Hyphomycetes may produce phialides solitarily (the genus Acremonium) or in loosely ordered structures (Trichoderma) or highly ordered structures (Aspergillus, Penicillium and related genera). Genera of interest here with less ordered phialidic structures can mostly be differentiated by macroscopic characters, e.g. colony diameters and colours. However, differentiating genera with highly ordered phialidic structures will necessitate careful microscopic examination. Phialides in Aspergillus, Penicillium and related genera are sometimes borne directly on a stalk or stipe which arises from a hypha; in other species, however, the phialides are borne from supporting cells, termed metulae (sing. metula) and in some species the metulae may in turn be supported by other cells, termed branches or rami (sing. ramus). The whole structure, including the stipe, is called a conidiophore. In Aspergillus, stipes are always robust, with thick walls and usually without septa; the stipe terminates in a more or less spherical swelling, the vesicle, which bears phialides, or metulae and phialides, over most of its surface. In Aspergillus, phialides (and metulae) are always produced simultaneously, and this feature can readily be recognised by examining young developing conidiophores (Fig. 8.1a). Similar structures, though smaller, are produced by some Penicillium species: these are clearly distinguished from Aspergillus species by stipes which are septate and by phialides which are produced over a period of time (successively; Fig. 8.2b). Most Penicillium species, and those of related genera, do not produce phialides on vesicles, but in a cluster directly on a stipe, or on metulae and/or rami. The fruiting structure in Penicillium and related genera is termed a penicillus, while that in Aspergillus (for want of a better term) is called a head (or more recently, an aspergillum, Klich, 2002). Coelomycetes. As noted earlier, Coelomycetes produce conidia within an enveloping body, termed a conidioma (pl. conidiomata). In Petri dish culture, conidiomata are produced on or just under the agar surface and are macroscopically visible, usually being 100-500 mm in diameter. Two kinds of conidioma are important here: the pycnidium, a more

5 Primary Keys and Miscellaneous Fungi

or less spherical body with one or more pores (ostioles) through which conidia are released, and the acervulus, a flat body from which conidia are released by lifting or rupturing of a lid. The majority of Coelomycetes are pathogens on plants and many have not been studied in pure culture. In consequence, their taxonomy is difficult and genera and species are often poorly delimited. For a complete account of Coelomycete taxonomy see Sutton (1980). Yeasts. Yeasts are fungi which have developed the ability to reproduce by forming single vegetative cells by budding or, in a few species, by fission, in a manner similar to bacteria. Like bacteria, and unlike fungal spores, such cells are metabolically active and may in turn reproduce by budding (or fission). Yeast cells may survive for long periods both in culture and in nature. In consequence many yeasts produce true spores rarely or not at all. Yeasts are readily distinguished from filamentous fungi on the agar plate by their soft-textured colonies and limited growth. They are usually also readily distinguished from bacteria by their raised and often hemispherical colonies, white or pink colours and lack of ”bacterial” odour. If in doubt, make a simple wet mount of a colony in water or lactofuchsin, add a cover slip and examine with the oil immersion lens. Yeast cells are larger than bacteria, measuring at least 3 2 mm and are nonuniform in size. If the culture is not too old, some cells will usually show developing buds. Yeasts cannot be classified solely by morphological features or growth on the standard media, and so are considered in a separate chapter (Chapter 10).

5.1 The General Key The taxonomic terms discussed above will enable the use of the general and miscellaneous keys which follow, although some other taxonomic terms may be introduced in discussions of particular genera. It is emphasised that these keys are designed for use on isolates which have been incubated for 7 days on the standard plating regimen outlined in Chapter 4. Colony diameters should be measured in millimetres from the reverse side by transmitted light. The general key has been designed to be as simple as possible and suitable for routine use, but it should be read in conjunction with the notes below it.

5.1 The General Key

55

General key to food spoilage fungi 1

No growth on any standard medium in 7 days Growth on one or more standard media

Chapter 9 – ”Xerophilic fungi” 2

2 (1)

Colonies yeasts, either recognisably so on isolation or in culture, i.e. colonies soft, not exceeding 10 mm diam on any standard medium Growth filamentous, exceeding 10 mm diam on one or more standard media

Chapter 10 – ”Yeasts” 3

3 (2)

Growth on CYA and/or MEA faster than on G25N Growth on G25N faster than on CYA and MEA

4 (3)

Hyphae frequently and conspicuously septate Hyphae lacking septa or septa rare

5 (4)

No mature spores psent in 7 days Mature spores psent in 7 days

6 9

6 (5)

Immature fruiting structures of some kind psent No fruiting structures (or spores) detectable by lowpower microscopy or wet mounts from CYA or MEA

7

5 Chapter 6 – ”Zygomycetes”

7 (6)

Colonies and fruiting structures white or brightly coloured Colonies or fruiting structures dark

8 (7)

Colonies and fruiting structures white Colonies or fruiting structures brightly coloured

9 (5)

Spores (conidia) less than 10 mm long, borne in chains on clustered fertile cells (phialides), on well-defined stipes Spores (conidia) of various sizes, borne singly or solitarily, or if borne in chains, then chains not in aggregates

10 (9)

11 (10)

Conidia blue or green, phialides produced successively on vesicles, vesicles less than 10 mm diam, stipes usually septate Conidia variously coloured, phialides and/or metulae produced simultaneously on vesicles, vesicles larger than 10 mm diam, stipes nonseptate

Couplet 1. No growth on any standard medium indicates an extreme xerophile, i.e. Xeromyces bisporus or a Chrysosporium species, or a nonviable culture. Next inoculate culture onto MY50G for 7 days at 258C. If growth occurs, enter the key in Chapter 9, ”Xerophilic Fungi”; no growth on MY50G indicates a nonviable culture. Chrysosporium and Xeromyces

See section on ”Miscellaneous fungi” below 8 Continue incubation; when spores mature, refer to section on ”Miscellaneous fungi” below Chapter 8 – ”Aspergillus and its teleomorphs” Chapter 7 – ”Penicillium and related genera”

Phialides or metulae and phialides borne on more or less spherical swellings on the stipe apices Phialides borne on penicilli, i.e. on unswollen stipes with or without intervening metulae and rami

5.1.1 Notes on the General Key

4 Chapter 9 – ”Xerophilic fungi”

10 See section on ”Miscellaneous fungi” below

11 Chapter 7 – ”Penicillium and related genera”

Chapter 7 – ”Penicillium and related genera”

Chapter 8 – ”Aspergillus and its teleomorphs”

isolates are usually white or rarely golden brown. If the original culture used as inoculum is coloured other than pure white or golden brown, it is probably nonviable. Couplet 2. Yeasts are usually readily distinguished by slow growth, soft, easily sampled colonies, small spherical to ellipsoidal cells, often of variable size and shape and by reproduction by budding. See Chapter 10, ”Yeasts” for identification procedures.

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5 Primary Keys and Miscellaneous Fungi

Couplet 3. The ability to grow more rapidly on G25N than on CYA or MEA indicates a xerophile (at least for keying purposes here). Check the key in Chapter 9, ”Xerophilic Fungi”. Some isolates can be identified from the standard plates, while others will require growth on CY20S or MY50G for identification. Couplet 4. The absence of septa in young, growing hyphae indicates an isolate belongs to subkingdom Zygomycotina, discussed here in Chapter 6 ”Zygomycetes”. Couplets 5, 6. Some isolates from a variety of fungal genera will not produce spores on the standard media in 7 days. Continue to incubate such cultures, pferably in diffuse daylight such as a laboratory window sill, at temperatures near 258C. Also inoculate such cultures onto two or three plates of DCMA and incubate these at 258C or thereabouts in darkness and in diffuse daylight or, if possible, under fluorescent illumination (see Chapter 4). After 1-2 weeks, check again for spores or fruiting bodies. If such structures are not seen, the isolate is unlikely to be significant in foods. Apparently asporogenous cultures should also be checked with a stereomicroscope while scraping up a sector of the colony with a needle. Fruiting bodies submerged in the agar will sometimes become visible with this technique. Couplet 7. Some isolates which produce white or brightly coloured fruiting bodies also produce very sparse aerial conidial structures which are easily overlooked. Check such cultures carefully with the stereomicroscope; if conidial structures are found, make a wet mount and reenter the key at Couplet 5.

A finely drawn glass needle will sometimes be of assistance in making mounts from delicate conidial structures on the colony. Nearly all dark fruiting structures encountered will mature at 258C within 2 weeks. Light does not usually influence this process. When mature spores are formed, refer to the following section.

5.2 Miscellaneous Fungi In this section are considered the genera which do not logically fit into some larger grouping considered elsewhere. Some are important in specific food spoilage problems, others are found in particular habitats such as cereals, while still others repsent the aerially dispersed fungal flora found as ubiquitous contaminants or saprophytes. As will be seen, they are a very heterogeneous collection. Most fungi significant in food spoilage or food contamination and not treated in other chapters are included here. It is inevitable, though, that occasional isolates from foods will not belong to the genera considered in this section. The key has not been designed to take account of this, as it would be a practical impossibility. So when an isolate appears to key out satisfactorily, it must be checked against the description to confirm the identification. Some isolates will of course belong to a recognisable genus, but not the species described; in that case the references indicated will provide further information. The miscellaneous fungal genera are considered in alphabetical order following the key.

Key to miscellaneous genera 1

Colonies on CYA and MEA not exceeding 60 mm diam in 7 days Colonies on CYA or MEA exceeding 60 mm diam in 7 days

2 12

2 (1)

Conidia borne within a fruiting body on or beneath the agar surface Conidia borne from aerial or surface hyphae

3 (2)

Mycelium and conidia hyaline or brightly coloured Mycelium and/or conidia dark coloured

4 11

4 (3)

Conidia with a single lateral septum Conidia nonseptate or with more than one septum

Trichothecium 5

5 (4)

Conidia borne from gradually tapering fertile cells (phialides) Conidia borne directly on hyphae or by budding or hyphal fragmentation

Phoma 3

6 7

5.2 Miscellaneous Fungi

6 (5)

Colonies exceeding 50 mm diam on CYA Colonies not exceeding 50 mm diam on CYA

7 (5)

Colonies exceeding 45 mm diam on MEA; conidia borne solely by the breakup of hyphae to form arthroconidia Colonies not exceeding 40 mm diam on MEA; conidia not exclusively arthroconidia

8 (7)

9 (8)

10 (9)

Budding hyphal fragments (10-50 mm long) psent Hyphal fragments absent; or if psent, not budding

11 (3)

Colonies low, mucoid and yeast-like, becoming grey to black in both obverse and reverse Colonies dry and velutinous, obverse green, reverse olive or deep blue black

57

Fusarium Acremonium

Geotrichum 8

Monascus 9

Endomyces 10 Hyphopichia Moniliella

Aureobasidium Cladosporium

12 (1)

Spores borne within an enclosed fruiting body on or under the agar surface Spores borne from aerial or surface hyphae

13 17

13 (12)

Spores consistently less than 15 mm long The larger or all spores more than 15 mm long

14 15

14 (13)

Fruiting bodies perithecia with stout, black hyphae attached to the walls Fruiting bodies pycnidia, without attached hyphae

15 (13)

Fruiting bodies roughly spherical (pycnidia) Fruiting bodies flat (acervuli)

16 (15)

Conidia hyaline or brightly coloured, nonseptate, without terminal appendages Conidia dark, with three or four septa and with spike-like, sometimes branched, terminal appendages

Chaetomium Phoma Lasiodiplodia 16 Colletotrichum Pestalotiopsis

17 (12)

Colonies and conidia hyaline or brightly coloured Colonies and/or conidia dark coloured

18 22

18 (17)

Colonies with grey or green areas Colonies white, orange, pink or purple

19 20

19 (18)

Colonies green Colonies grey

20 (18)

Colonies low and persistently white Colonies floccose, white or becoming brightly coloured

Geotrichum 21

21 (20)

Colonies pdominantly orange, orange conidia shed profusely around the Petri dish rim Colonies white, pink or purple, sporulation on MEA weak or absent, better on DCPA under lights

Chrysonilia

22 (17)

Conidia consistently less than 15 mm long Conidia frequently exceeding 15 mm long

Trichoderma Botrytis

Fusarium 23 25

58

23 (22)

5 Primary Keys and Miscellaneous Fungi

Conidiophores long, branched, apically swollen, bearing closely packed pale brown conidia Conidiophores short or ill-defined, dark brown or black conidia borne irregularly

Botrytis 24

24 (23)

Conidia dark brown, often with a lighter coloured band around the periphery Conidia uniformly jet black

Arthrinium Nigrospora

25 (22)

Conidia approximately spherical Conidia elongate

Epicoccum 26

26 (25)

Conidia with transverse septa (or thick walls between cells) only Larger conidia with both transverse and longitudinal septa or irregularly septate

27 30

27 (26)

Conidia clavate (club shaped), often with long hyphal appendages at the apices; found almost exclusively on rice Conidia cylindroidal, ellipsoidal or an elongate ”D” shape; source more general

Trichoconiella 28

28 (27)

Conidia cylindroidal, with parallel sides except at the terminal cells Conidia fusoid, narrowing from the central cells to the terminal cells, often bent or ”D” shaped

Drechslera

29 (28)

Conidia not exceeding 40 mm long Conidia usually exceeding 40 mm long

Curvularia Bipolaris

30 (26)

Conidia clavate (club shaped) Conidia spherical to roughly ellipsoidal or short cylindrical

Alternaria 31

31 (30)

Conidia often tapering towards the base and sometimes pointed, i.e. pyriform or apiculate Conidia spherical to short cylindrical, not tapering from base to apex, with rounded ends

5.3 Genus Acremonium Link Commonly referred to as Cephalosporium Corda in p-1970 literature, Acremonium is a large and varied genus characterised by the production of small, hyaline, single-celled conidia borne singly, i.e. successively but not connected to each other, from solitary phialides. A variety of species have been recorded from foods from time to time. One species, A. strictum, described here, is of relatively common occurrence. In this species, the phialides gradually taper to the apices without basal thickening or formation of a distinct neck, and conidia aggregate in balls of slime. Under the stereomicroscope, the slime balls look like large single spores, but their true nature becomes evident in wet mounts. A. strictum appears mainly in the earlier food literature under the name C. acremonium. As noted by Domsch et al. (1980), this name has been used for a variety of species, so that reports on physiology and occurrence are unreliable (Fig. 5.1).

Acremonium strictum W. Gams

29

Ulocladium Stemphylium

Fig. 5.1

Cephalosporium acremonium (name of uncertain application; no valid authority).

Colonies on CYA 20-30 mm diam, white or orange to pink, dense to floccose or funiculose; reverse pale or with orange to pink tones. Colonies on MEA 13-20 mm diam, similar to those on CYA or of slimy texture. Colonies on G25N 1 mg/kg, but can occur at lower levels sometimes. In ppubertal gilts, clinical signs include vulval swelling, uterine enlargement and mammary development. Mature sows can develop ovarian atrophy, constant oestrus and pseudopgnancy (Hagler et al., 2001). Prepubertal male pigs can undergo a feminising effect with mammary development, decreased testicular size and loss of libido, but mature boars are resistant (Hagler et al., 2001). It was once claimed that humans had shown similar signs in Puerto Rico, but an FDA investigation failed to confirm this (Goodman et al., 1987). T-2 toxin. T-2 toxin is the cause of alimentary toxic aleukia (ATA) a devastating disease which occurred in the USSR during and after Second World War, in times of extreme food shortage resulting in consumption of overwintered cereals (Joffe, 1978; Beardall and Miller, 1994). Many people, probably hundreds of thousands, died as a result (Marasas et al., 1984). ATA was characterised by leucopoenia, bleeding from nose, throat and gums, haemorrhagic rash, exhaustion of the bone marrow and fever. Vomiting, nausea, diarrhoea and abdominal pain also usually occurred. Decrease in immunological functions led to susceptibility to bacterial and viral diseases, and often death (Joffe, 1978; Beardall and Miller, 1994). Haemorrhagic syndrome in cattle, pigs and poultry in the United States in the 1960s was also probably due to T-2 toxin (Desjardins, 2006). T-2 toxin is produced by F. sporotrichioides and, less commonly, F. poae (Desjardins, 2006). It appears to be produced only under cold conditions, and fortunately is now uncommon. The occurrence, toxicity and biology of T-2 toxin were examined in detail by JECFA (2001). They established that T-2 (and its metabolite HT-2) were immunotoxic and haemotoxic compounds in several animal species after short-term intake. Long-term effects could not be evaluated. T-2 was at most weakly genotoxic. In the absence of long-term studies, T-2 was not classifiable as to carcinogenicity (JECFA, 2001). Fumonisins. Fumonisins are produced by F. verticillioides (formerly known as F. moniliforme), by the closely related species F. proliferatum, uncommonly by F. subglutinans and F. oxysporum,

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5 Primary Keys and Miscellaneous Fungi

with neural tube defects such as spinal bifida in a population along the Texas-Mexican border (Desjardins, 2006). For the human population, a provisional maximum tolerable daily intake (PMTDI) of 2 mg/kg body weight per day was established by JECFA (2001). Determining which particular Fusarium species produces which mycotoxins has not been easy. Unstable taxonomy and misidentification have combined to cloud the picture. In a major contribution, Marasas et al. (1984) examined several hundred Fusarium isolates, identifying them according to Nelson et al. (1983) and then assessing mycotoxin production. Sections on mycotoxins in the species descriptions which follow are based on Marasas et al. (1984), Desjardins (2006) and Leslie and Summerell (2006). Claims concerning mycotoxin production by particular species in more recent references should be treated with some caution. Cultural instability. Many Fusarium species are notorious for their instability in culture. Isolates of some species will degenerate quickly, often after only one or two transfers. For this reason, it is important to identify Fusaria as soon as possible after primary isolation. Pure cultures for identification are traditionally started from a single germinated spore, as the mass transfer of Fusaria appears to increase the rate of deterioration of strains in culture. Identification procedures. Identification of Fusarium isolates is often difficult, but the task can be made easier by observing a few basic rules:

identify cultures as soon as possible after primary isolation;

always grow cultures for identification from single germinated conidia; standardised media and incubation conditions; use a light bank (Chapter 4) whenever possible.

use

Diagnostic features. The main characters used to distinguish species of Fusarium are (1) the size and shape of the macroconidia; (2) the psence or absence of microconidia; (3) the manner in which microconidia are produced; (4) the type of phialide on which microconidia are produced; (5) the psence or absence of chlamydoconidia; and (6) the colours and morphology of colonies on PDA.

5.17 Genus Fusarium

93

The morphology of macroconidia is a principal diagnostic feature for Fusarium species. Macroconidia generally have at least three septa, with a differentiated apical cell which may be pointed, rounded, hooked or filamentous, and a basal cell which may be foot-shaped, with a distinct heel, or just slightly notched. Fusarium macroconidia generally exhibit some degree of curvature, the convex and concave sides being referred to as the dorsal and ventral sides, respectively. Although some macroconidia are usually produced in the aerial mycelium, the shape and size of those in sporodochia are more regular and are used for identification purposes where possible. Microconidia are usually produced in the aerial mycelium and their shape can be very important in Fusarium identification. Most species which produce microconidia form only a single type, the most common shape being ellipsoidal to clavate. However, F. poae produces spherical to apiculate microconidia and F. sporotrichioides produces a variety of shapes: ellipsoidal, pyriform and spherical. The method of production of microconidia and the types of phialides on which they are borne are also useful diagnostic criteria. F. verticillioides produces its microconidia in long, delicate, dry chains, which are best observed by using the 10 objective of the

Key to Fusarium species included here 1

Microconidia abundant Microconidia rare or absent

2 9

2 (1)

Colonies on PDA with mycelium and/or reverse coloured greyish rose or burgundy Colonies on PDA in shades of cream, pale salmon or violet

3 5

3 (2)

4 (3)

5 (2)

6 (5)

F. poae 4

F. chlamydosporum F. sporotrichioides

6 7 F. verticillioides F. proliferatum

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5 Primary Keys and Miscellaneous Fungi

7 (5)

Colonies cream or bluish, sporodochia cream Colonies pale salmon or violet, sporodochia salmon

8 (7)

Microconidia borne on short, stout monophialides; chlamydoconidia usually produced Microconidia borne on polyphialides and slender monophialides; chlamydoconidia not produced

9 (1)

Colonies cream, pale salmon or brown Colonies greyish rose to burgundy

10 (9)

Macroconidia cigar- or spindle-shaped, produced in the aerial mycelium Macroconidia obviously curved, produced in sporodochia

11 (9)

Macroconidia robust, ventral side straight; aerial mycelium tan to brown Macroconidia delicate and slender, slightly or definitely curved; aerial mycelium white or pinkish

12 (11)

Macroconidia longer and narrower, maximum width 5.5 mm 13 (11)

Macroconidia with elongated basal cell and long, whip-like apical cell Macroconidia with basal and apical cells not obviously elongated

14 (13)

Macroconidia delicate and needle-like, with sides almost parallel Macroconidia with slight to definite curvature

Telemorph: Gibberella acuminata Wollenw. Colonies on CYA 40-50 mm diam, of dense, felty mycelium, white to greyish rose or greyish magenta; reverse uniformly pale or with areas of greyish rose. Colonies on MEA 45-65 mm diam, yellow brown centrally, greyish rose at the margins; reverse deep brownish yellow to brownish orange, occasionally pale. Colonies on G25N 9-15 mm diam. At 58C, colonies 7-12 mm diam. No growth at 378C. On PDA, colonies usually covering the whole Petri dish, of dense to floccose white to pale salmon mycelium, sometimes greyish rose at the margins; reverse dark ruby centrally, greyish ruby at the margins. On DCPA, colonies sparse, of floccose to funiculose white to pale salmon mycelium; reverse pale or with brownish red annular rings. Macroconidia relatively slender, usually with five septa, but three and four septa not uncommon, with a long, tapering apical cell and foot-shaped basal cell, distinctly but not highly curved, with

F. oxysporum F. subglutinans 10 11

Macroconidia short and stout, up to 7 mm wide

Fusarium acuminatum Ellis & Everh. Fig. 5.22

F. solani 8

F. semitectum F. equiseti 12 13 F. culmorum (See F. graminearum) F. graminearum F. longipes 14 F. avenaceum F. acuminatum

the widest point often one third of the distance from the base, giving a ”bottom-heavy” appearance; microconidia produced sparsely by some isolates; chlamydoconidia produced, relatively slowly. Distinctive features. Ruby to dark ruby reverse colours on PDA, and relatively slender, slightly curved macroconidia, usually with five septa, are the distinctive features of Fusarium acuminatum. However, unless chlamydoconidia are psent, this species can be confused with F. avenaceum (see below). F. armeniacum is very closely related (Burgess and Summerell, 2000). Taxonomy. Perithecia of Gibberella acuminata Wollenw., the teleomorph of F. acuminatum, are formed in the laboratory when opposite mating types are inoculated onto sterile wheat straws (Booth, 1971). Isolates of F. acuminatum show considerable variability in culture, and this variability, correlated with secondary metabolite production (Logrieco et al., 1992), has been shown to be due to the fact that two species were included within F. acuminatum. F. armeniacum (Forbes et al.) L.W. Burgess and Summerell, pviously described as F. acuminatum var. armeniacum G.A.

5.17 Genus Fusarium

95

Fig. 5.22 Fusarium acuminatum (a) colonies on PDA and DCPA, 7 d, 258C; (b, c) macroconidia, bar = 10 mm

Forbes et al. has been elevated to species level (Burgess and Summerell, 2000) being distinguished on morphological grounds, isoenzyme patterns, molecular markers and mycotoxin profiles. Macroconidia of F. armeniacum are produced in distinct apricot coloured sporodochia. Physiology. Some isolates of Fusarium acuminatum have antioxidant enzyme activity (Kayali and Tarhan, 2005). Mycotoxins. Most of the mycotoxin production reported from Fusarium acuminatum, particularly the production of trichothecene toxins, is probably more correctly due to F. armeniacum (Desjardins, 2006; Leslie and Summerell, 2006). Isolates producing type A trichothecenes including T-2 toxin, HT-2 toxin and diacetoxyscirpenol (see Pitt and Hocking, 1997) are most likely to be F. armeniacum. In a survey of 25 isolates of F. acuminatum from different sources and geographic locations,

Logrieco et al. (1992) pided the isolates into three categories, (a) enniatin B and/or moniliformin producers (probably F. acuminatum sensu stricto), (b) T-2, HT-2 and/or neosolaniol producers (probably F. armeniacum) and (c) nontoxigenic. F. acuminatum sensu stricto produces moniliformin (Chelkowski et al., 1990; Logrieco et al., 1992) and enniatins (Logrieco et al., 1992; Kononeko et al., 1993; Desjardins, 2006; Leslie and Summerell, 2006) as well as some other minor toxins (Desjardins, 2006). Ecology. Fusarium acuminatum has been isolated from a wide variety of plants throughout the world. Although some isolates may cause severe root rot in particular legume species (Leslie and Summerell, 2006), F. acuminatum is generally regarded as a saprophyte. It has been reported to cause rot in pumpkins (Elmer, 1996), is one cause of rot in stored potatoes (Theron and Holz, 1990) and is weakly pathogenic in

96

5 Primary Keys and Miscellaneous Fungi

bananas (Jime´nez et al., 1993). It is quite common in poor quality wheat from cool temperate zones (Mills and Wallace, 1979; Abramson et al., 1987). It has been isolated from developing peanut pods (Barnes, 1971), barley (Abdel-Kader et al., 1979) and, in our laboratory, from rain damaged sorghum and soybeans. The incidence of F. acuminatum in tropical commodities was low (Pitt et al., 1993, 1994). References. Domsch et al. (1980), under G. acuminata; Nelson et al. (1983); Desjardins (2006); Leslie and Summerell (2006).

Fusarium avenaceum (Fr.) Sacc.

Fig. 5.23

Teleomorph: Gibberella avenacea R.J. Cooke Colonies on CYA covering the whole Petri dish, moderately deep to deep, of open, floccose

mycelium coloured white, very pale rose or deeper greyish rose; reverse varying from pale to pale yellow or with areas of greyish rose or sometimes uniformly deep burgundy. Colonies on MEA 45- 55 mm diam, low to moderately deep, of open floccose to funiculose mycelium, coloured white, pale rose or greyish rose, sometimes brown centrally; reverse brownish orange, sometimes paler centrally or at the margins. Colonies on G25N 9- 15 mm diam. At 58C, colonies 10-12 mm diam. No growth at 378C. On PDA, colonies moderately deep to deep, of dense mycelium coloured white, pale salmon or sometimes dark brownish red, with central masses of reddish orange sporodochia, sometimes surrounded by an outer ring of paler sporodochia; reverse greyish red, with darker annular rings, paler towards the margins. On DCPA, colonies deep, of moderately dense white to pale salmon mycelium with a central mass of orange to

Fig. 5.23 Fusarium avenaceum (a) colonies on PDA and DCPA, 7 d, 258C; (b, c) macroconidia, bar = 10 mm

5.17 Genus Fusarium

salmon sporodochia, often surrounded by concentric rings of sporodochia; reverse pale. Macroconidia long, slender, with four to seven septa, thin-walled, straight or slightly curved, with a tapering apical cell and a notched or foot-shaped basal cell; microconidia produced sparsely by some isolates; chlamydoconidia absent. Distinctive features. Fusarium avenaceum is distinguished by thin walled, needle-like macroconidia and by the absence of chlamydoconidia. Despite the fact that F. avenaceum and F. acuminatum are not considered by Fusarium taxonomists to be closely related, these two species can be difficult to distinguish, as isolates with macroconidia of intermediate form are not uncommon. Colony diameters on PDA at 308C after 3 days can be a useful differentiating feature: under these conditions colonies of F. avenaceum are usually 8- 15 mm diam, whereas those of F. acuminatum are 15-28 mm diam (Burgess et al., 1994). Isolates of F. avenaceum show an unusually broad range of colours on PDA and also have a very broad host range. However, extensive genetic analysis has shown no bases from splitting the species, and pathogenicity tests on single strains have confirmed the broad host range (Nalim, 2004). Taxonomy. The teleomorph of Fusarium avenaceum is Gibberella avenacea R.J. Cook. It is not usually seen in culture on the media used here. Physiology. The optimum growth temperature for Fusarium avenaceum is 358C, the minimum near -38C and the maximum 318C (Domsch et al., 1980). The minimum aw for growth is approximately 0.90 at 258C (Magan and Lacey, 1984c), and the pH optimum ranges between 5.4 and 6.7 (Domsch et al., 1980). Mycotoxins. This species has been reported to produce a variety of trichothecene and other mycotoxins. However, Nelson et al. (1983) regarded only reports of moniliformin production as accurate. Later reports have confirmed this (Chelkowski et al., 1990; Abbas et al., 1991; Bosch and Mirocha, 1992). Reports of production of fusarin C (Farber and Sanders, 1986; Thrane, 1988; Leonov et al., 1993) and enniatins (Blais et al., 1992; Kononeko et al., 1993) also appear to be reliable (Desjardins, 2006; Leslie and Summerell, 2006). Production of any trichothecene toxin has not been confirmed, and Fusarium avenaceum does not carry the tri5 gene which is essential for trichothecene production (Tan and Niessen, 2003).

97

Ecology. Fusarium avenaceum has a worldwide distribution wherever crops are grown, but is relatively uncommon in food commodities. It is a major component of Fusarium head blight in cereals in Europe, the US Pacific Northwest region and Canada (Desjardins, 2006). Logrieco et al. (2002) identified F. avenaceum as a component of Fusarium ear rot of maize in Europe. It has been reported from barley (Flannigan, 1969; Petters et al., 1988; Stenwig and Liven, 1988) where it may inhibit germination of malting grains (Hudec, 2007), but is of minor importance in gushing of beer (Niessen et al., 1992). Other reported sources are sorghum (Onyike and Nelson, 1992), peanuts (Joffe, 1969), pigeon peas (Maximay et al., 1992) and, in our laboratory, triticale. F. avenaceum has been reported to cause spoilage of cool-stored broccoli (Mercier et al., 1991), dry rot of stored carrots in Italy (Marziano et al., 1992) and dry rot of rutabaga (swede turnip) in Canada (Peters et al., 2007). It has occasionally caused spoilage of apples, pears, asparagus, tomatoes, eggplant and potatoes (Snowdon, 1990, 1991) and has been reported as a postharvest pathogen of stonefruit in New Zealand (Hartill and Broadhurst, 1989). References. Domsch et al. (1980), as G. avenacea; Nelson et al. (1983); Leslie and Summerell (2006).

Fusarium chlamydosporum Wollenw. & Reinking Fig. 5.24 Fusarium fusarioides (Gonz. Frag. & Cif.) Booth

Colonies on CYA covering the whole Petri dish, of low to moderately deep floccose mycelium, coloured white to pale rosy pink, often with surface appearing powdery due to production of microconidia; reverse pale to greyish rose or brownish red. Colonies on MEA 55-70 mm diam, of low, moderately dense mycelium in shades of yellow brown, or greyish rose to greyish ruby, paler at the margins; reverse deep yellow brown to orange brown. Colonies on G25N 15-20 mm diam. At 58C, colonies 1-2 mm diam. At 378C, colonies 5-15 mm diam. On PDA, colonies of felty mycelium, coloured pale salmon, sometimes browner, or with patches of greyish red, often with a powdery appearance from profuse microconidial production; reverse deep violet brown to dark ruby, paler at the margins. On DCPA, colonies of sparse, floccose, pale salmon mycelium, often powdery with microconidia, showing poorly defined annulations; macroconidia

98

5 Primary Keys and Miscellaneous Fungi

Fig. 5.24 Fusarium chlamydosporum (a) colonies on PDA and DCPA, 7 d, 258C; (b, c) polyphialides, bar = 10 mm; (d) macroconidia, bar = 10 mm; (e) microconidia, bar = 10 mm

occasionally produced near the colony centres in salmon sporodochia; reverse pale. Macroconidia often rare, relatively short and stout, usually with three to five septa, slightly curved; microconidia produced abundantly from polyphialides in the aerial mycelium, with zero to two septa, fusiform to slightly clavate. Chlamydoconidia usually abundant in older cultures, produced singly, in pairs or in clumps. Distinctive features. The psence of abundant fusiform microconidia borne on polyphialides is the most outstanding feature of Fusarium chlamydosporum. Also colonies on PDA have dark violet brown to dark ruby reverse colours.

Taxonomy. Fusarium chlamydosporum has priority over F. fusarioides as the correct name for this species (Domsch et al., 1980; Nelson et al., 1983; Leslie and Summerell, 2006). Physiology. This species has minimum, optimum and maximum temperatures for growth of 5, 27 and 378C (Seemu¨ller, 1968). Mycotoxins. Production of type A trichothecenes (including T-2 toxin, HT-2 toxin, monoacetoxyscirpenol, neosolaniol and iso-neosolaniol) by Fusarium chlamydosporum was reported by Park and Chu (1993); however, subsequent studies have found no evidence of trichothecene production in this species (Desjardins, 2006). Moniliformin is the

5.17 Genus Fusarium

major mycotoxin produced by F. chlamydosporum (Marasas et al., 1984; Desjardins, 2006). Ecology. Fusarium chlamydosporum is mainly an inhabitant of soils in warmer climates (Domsch et al., 1980; Leslie and Summerell, 2006), and is not regarded as a plant pathogen or spoilage fungus. However, it is commonly isolated from grains in drier areas, particularly in the Middle East, southern Europe, central Asia and Australia (Leslie and Summerell, 2006), and has also been isolated from pearl millet (Wilson et al., 1993; Jurjevic et al., 2007), pecans (Huang and Hanlin, 1975) and sorghum (Rabie et al., 1975; Onyike and Nelson, 1992). A low incidence of F. chlamydosporum was found in peanuts from both Indonesia and the Philippines (Pitt et al., 1998a) and from mung beans and sorghum in Thailand (Pitt et al., 1994). Involvement in dry rot of potatoes has also been reported (Somani, 2004; Esfahani, 2006). References. Domsch et al. (1980); Nelson et al. (1983); Leslie and Summerell (2006).

99

Fusarium culmorum (W.G. Smith) Sacc. Fig. 5.25 Colonies on CYA covering the whole Petri dish, of dense felty mycelium, often with a floccose overlay, sometimes reaching the Petri dish rim, pale red to pastel red; reverse pastel red to deep red. Colonies on MEA 60 mm or more diam, floccose, in age often reaching the Petri dish lid, pale red to pastel red, commonly with a greyish orange to yellowish brown overlay; reverse brown to reddish brown. Colonies on G25N usually 5-10 mm diam, mycelium orange white, reverse yellow to orange. At 58C, germination. No growth at 378C. On PDA, colonies covering the whole Petri dish, of dense to floccose mycelium, pale red and pale yellow brown; reverse red to deep red. On DCPA, colonies 50-65 mm diam, of sparse mycelium, orange to pinkish white, bearing abundant macroconidia in orange sporodochia; reverse dull orange brown.

Fig. 5.25 Fusarium culmorum (a) colonies on PDA and DCPA, 7 d, 258C; (b, c) macroconidia, bar = 10 mm

100

Macroconidia relatively short, wide and only slightly curved, with four to five septa, 30-45 mm long, with rounded or sometimes papillate apical cells; basal cells with a slight to definite notch, sometimes papillate. Microconidia not produced. Chlamydoconidia sometimes formed, in conidia, or intercalary in the hyphae, singly or in chains, 9-14 mm diam, smooth walled. Distinctive features. Short, stout macroconidia are the prime feature distinguishing Fusarium culmorum from most other species. F. culmorum may be confused with F. crookwellense L.W. Burgess et al., but macroconidia of the latter species have a distinctly foot-shaped basal cell, whereas those of F. culmorum are shorter and stouter, and the basal cell is not distinctly foot-shaped. Taxonomy. No teleomorph is known for this species. Physiology. Fusarium culmorum has been reported to be psychrotrophic, growing down to 08C, with an optimum at 218C and a maximum of only 318C (Arsvoll, 1975); however, Magan and Lacey (1984c) reported growth at 358C. The minimum aw for growth is 0.87 at 20-258C and pH 6.5: at pH 4.0, growth did not occur below 0.90 aw (Magan and Lacey, 1984a). A strain of F. culmorum produced deoxynivalenol optimally at 258C, but only between 0.995 and 0.97-0.96 aw (Hope and Magan, 2003; Hope et al., 2005). At 158C, deoxynivalenol was produced in lower concentrations later in the growth cycle, but over a slightly greater aw range (0.995 to 0.95-0.94 aw). The dynamics of nivalenol production for this strain was similar (Hope and Magan, 2003). Zearalenone production by F. culmorum was reported to be optimal above 258C (Bottalico et al., 1982). F. culmorum is very tolerant of low O2 tensions (Magan and Lacey, 1984b). Radiation resistance of F. culmorum was relatively high: up to 0.8 kGy were needed for a tenfold reduction in spore numbers on grain, and up to 1.39 kGy on media (O’Neill et al., 1991). Mycotoxins. Fusarium culmorum produces a variety of mycotoxins: indeed the list in our files is of more than 40 compounds. However, the most important toxins confirmed to be produced by this species (Nelson et al., 1983; Marasas et al., 1984; Desjardins, 2006; Leslie and Summerell, 2006) are deoxynivalenol, nivalenol and their derivatives (Abramson et al., 2001; Hestbjerg et al., 2002;

5 Primary Keys and Miscellaneous Fungi

Chandler et al., 2003; Jennings et al., 2004) and zearalenone (Bakan et al., 2001; Hestbjerg et al., 2002; Llorens et al., 2004a; Brinkmeyer et al., 2005). Moniliformin production was reported by Scott et al. (1987) but was not detected in 42 isolates of F. culmorum from Canada by Abramson et al. (2001). Reports of production of type A trichothecenes (T-2 toxin, HT-2 toxin) have not been substantiated (Leslie and Summerell, 2006). The existence of two chemotypes of Fusarium culmorum, those that produce deoxynivalenol and those that produce nivalenol (Miller et al., 1991), has been confirmed by molecular studies. Within the trichothecene gene cluster, isolates possessing the Tri7 and Tri13 genes produce nivalenol and related compounds, whereas sequences in the Tri3, Tri5 and Tri6 genes are associated with deoxynivalenol production (Chandler et al., 2003; Jennings et al., 2004; Quarta et al., 2005, 2006). Of 55 European isolates examined by Quarta et al. (2005), 11 were the nivalenol chemotype, and the remainder were deoxynivalenol producers. Jennings et al. (2004) examined 153 isolates from England and Wales, and found that the DON chemotype was dominant over the NIV chemotype (59% vs 41%, respectively). Lauren et al. (1992) examined 45 isolates of F. culmorum from New Zealand soil and pasture and found none produced deoxynivalenol or its monoacetyl isomers. Two chemotypes were identified, one producing diacetylnivalenol with culmorin as the major metabolite accounted for 95% of isolates, while the other chemotype produced diacetoxyscirpenol. Ecology. This species has a worldwide distribution in soil and as a pathogen of cereals and other hosts, with a higher incidence in temperate climates (Domsch et al., 1980; Nelson et al., 1983; Leslie and Summerell, 2006). It is an important component of the cohort of Fusarium species that cause head blight of wheat and associated cereal crops in Europe, Canada, China and other areas with cool weather during the growing season (Desjardins, 2006). In wheat it causes extensive internal damage to the grain, and reductions in flour yield and baking quality (Meyer et al., 1986). F. culmorum was reported as the dominant Fusarium species on barley in Europe (see Pitt and Hocking, 1997). It also occurs in triticale (Perkowski et al., 1988). F. culmorum has been identified as a component of Fusarium ear rot of maize in Europe (Logrieco

5.17 Genus Fusarium

101

et al., 2002). F. culmorum was one cause of crown rot in bananas (Wade et al., 1993), and is a minor cause of spoilage of apples and pears (Snowdon, 1990). References. Domsch et al. (1980); Nelson et al. (1983); Desjardins (2006); Leslie and Summerell (2006).

Fusarium equiseti (Corda) Sacc.

Fig. 5.26

Teleomorph: Gibberella intricans Wollenw. Colonies on CYA filling the whole Petri dish, often to the lid, of dense to floccose white mycelium; reverse pale or pale salmon. Colonies on MEA

covering the whole Petri dish, of open, floccose white to pale brown mycelium; reverse pale, or sometimes showing areas of pale greyish red. Colonies on G25N 12-20 mm diam. At 58C, colonies of 1- 4 mm diam produced. At 378C, usually no growth, although in isolates from the tropics, colonies up to 35 diam Produced. On PDA, colonies of dense to floccose mycelium, white to pale salmon, becoming brown with age, with a central mass of orange to brown sporodochia, sometimes surrounded by poorly defined sporodochial rings; reverse pale salmon, often with a brown central area and brown flecks. On DCPA, colony appearance usually dominated by salmon, orange or brownish

Fig. 5.26 Fusarium equiseti (a) colonies on PDA and DCPA, 7 d, 258C; (b, c) macroconidia, bar ¼ 10 mm; (d) chlamydoconidia, bar ¼ 10 mm

102

421

The factors enabling fungi to cause spoilage in cheese are the ability to grow at refrigeration temperatures, to grow in low oxygen concentrations, lipolytic activity, resistance to the pservative action of free fatty acids and growth at reduced aw. Penicillium roqueforti and P. commune meet all these criteria and are thus the most successful spoilage moulds on cheese. Toxin production (roquefortine and PR toxin from P. roqueforti and cyclopiazonic acid from P. commune) is a definite, though probably small, hazard. PR-imine was detected in 50 of 60 samples of blue-vein cheese, but PR toxin was not (Siemens and Zawistowski, 1993). Roquefortine was detected in 1 of 10 samples of Valdeon cheese, a blue´ mould ripened Spanish variety (Lopez-Dias et al., 1996), in all 11 samples of European blue-mould cheeses purchased in Finland (Kokkonen et al., 2005a) and in all 30 samples of European bluemould cheese examined in Italy (Finoli et al., 2001). The levels of roquefortine detected by Kokkonen et al. (2005b) ranged from 0.8 to 12 mg/kg, whereas levels reported by Finoli et al. (2001) were lower (0.08-1.47 mg/kg). Mycophenolic acid was detected in 4 of 12 samples of mouldy Spanish Manchego ´ cheese (Lopez-Dias et al., 1996) and 1 of 11 samples of blue-mould cheese in Finland (Kokkonen et al., 2005a). Ochratoxin A has recently been reported by Dall’Asta et al. (2008) in blue-mould cheeses from Italy (23 of 54 samples) and France (7 of 14 samples) at levels from 0.25 to 3.0 mg/kg. Cyclopiazonic acid detected in six samples of Italian Taleggio, a soft, smear-ripened cheese, was confined mainly to the rind (Finoli et al., 1999). Sterigmatocystin produced by Aspergillus versicolor was detected in the surface layer of hard cheeses in the Netherlands (Northolt et al., 1980). Mycotoxin levels reported in cheese are not usually considered to be of public health significance. Mouldy cheese is unsuitable for sale and for manufacturing purposes. Protection from the Penicillia relies on clean production conditions, low temperature storage, low oxygen atmospheres, integrity of packaging materials, intact rinds, pservative impgnated wrappers and rapid turnover of stock.

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Media Appendix

Aspergillus flavus and parasiticus agar (AFPA) Peptone, bacteriological Yeast extract Ferric ammonium citrate Chloramphenicol Agar Dichloran (0.2% in ethanol, 1.0 ml) Water, distilled

10 g 20 g 0.5 g 100 mg 15 g 2 mg 1l

After addition of all ingredients, sterilise by autoclaving at 1218C for 15 min. The final pH of this medium is 6.0-6.5. Creatine sucrose neutral agar (CSN) CS concentrate Sucrose Creatine KH2PO4 Bromocresol purple Agar Water, distilled to

10 ml 10 g 5.0 g 1.0 g 0.05 g 15 g 1l

Creatine sucrose (CS) concentrate KCl MgSO4 7H2O FeSO4 7H2O ZnSO4 7H2O CuSO4 5H2O Water, distilled to

5g 5g 0.1 g 0.1 g 0.05 g 100 ml

Sterilise by autoclaving at 1218C for 15 min. Final unadjusted pH is approximately 6.8. A pH between 5.5 and 6.8 is satisfactory. For identification of Penicillium subgenus Penicillium species. Czapek concentrate NaNO3 KCl

30 g 5g

MgSO4 7H2O FeSO4 7H2O Water, distilled

5g 0.1 g 100 ml

Czapek concentrate will keep indefinitely without sterilisation. The pcipitate of Fe(OH)3 which forms in time can be resuspended by shaking before use. Czapek iprodione dichloran agar (CZID) Sucrose Yeast extract Chloramphenicol Dichloran (0.2% in ethanol, 1 ml) Czapek concentrate Trace metal solution Agar Water, distilled Iprodione (suspension)

30 g 5g 100 mg 2 mg 10 ml 1 ml 15 g 1l 1 ml

Add iprodione suspension . Kvashnina, E.S. 1976. (Physiological and ecological characteristics of Fusarium species Sect. Sporotrichiella). Mikol. Fitopatol. 10: 275-281. Kwasna, H., Ward, E. and Bateman, G.L. 2006. Phylogenetic relationships among Zygomycetes from soil based on ITS1/2 rDNA sequences. Mycol. Res. 110: 501-510. La Guerche, S., Dauphin, B., Pons, M., Blancard, D. and Darriet, P. 2006. Characterization of some mushroom and earthy off-odors microbially induced by the development of rot on grapes. J. Agric. Food Chem. 54: 9193-9200. Labuda, R. and Tancˇinova´, D. 2003. Eupenicillium ochrosalmoneum, a rare species isolated from a feed mixture in Slovakia. Biologia, Bratislava 58: 1123-1126. Lacey, J. 1980. Colonization of damp organic substrates and spontaneous heating. In Microbial Growth and Survival in Extreme Environments, eds G.W. Gould and J.E.L. Corry. London: Academic Press. pp. 53-70. Lahali, R., Serrhini, M.N. and Jijakli, M.H. 2004. Efficacy assessment of Candida oleophila (strain O) and Pichia anomala (strain K) against major postharvest diseases of citrus fruits in Morocco. Commun. Appl. Biol. Sci. Ghent Univ. 69: 601-609. Lahali, R., Serrhini, M.N. and Jijakli, M.H. 2005. Studying and modelling the combined effect of temperature and water activity on the growth rate of P. expansum. Int. J. Food Microbiol. 103: 315-322. Lahlali, R., Serrhini, M.N., Friel, D. and Jijakli, M.H. 2007. Predictive modelling of temperature and water activity (solutes) on the in vitro radial growth of Botrytis cinerea Pers. Int. J. Food Microbiol. 114: 1-9. Laich, F., Fierro, F., Cardoza, R.E. and Martin, J.F. 1999. Organization of the gene cluster for biosynthesis of penicillin in Penicillium nalgiovense and antibiotic production in cured dry sausages. Appl. Environ. Microbiol., 65: 1236-1240. Lamper, C., Teren, J., Bartok, T., Komoroczy, R., Mesterhazy, A. and Sagi, F. 2000. Predicting DON contamination in Fusarium-infected wheat grains via determination of the ergosterol content. Cereal Res. Commun. 28: 337-344. Lanciotti, R., Sinigaglia, M., Gardini, F. and Guerzoni, M.E. 1998. Hansenula anomala as spoilage agent of cream-filled cakes. Microbiol. Res. 153: 145-148.

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Index

Note: Bold face denotes fungal species description A Absidia corymbifera, 148 in tree nuts, 408 oxygen requirements, 7 ramosa, 149 Acetic acid pserves, spoilage of, 419 Candida krusei, 363 Moniliella acetoabutans, 130 Pichia membranaefaciens, 371 Acremonium, 57, 58 strictum, 58 Acute cardiac beriberi, 197 Aflatoxins, 307-309 from Aspergillus flavus, 307 from Aspergillus nomius, 311 from Aspergillus parasiticus, 322 coconut cream agar for detection of, 28 from milk, 308 Air sampling, 23 Alimentary toxic aleukia (ATA), 1, 91, 109, 117 Altenuene, 62 Alternaria, 58, 60, 142 alternata, 60, 63 in cantaloupes, 387 in cereals, 395, 396 in citrus, 385 in figs, 389 in mangoes, 390 mycotoxins, 61 in pome fruits, 386 in rice, 396 in sorghum, 398 in soybeans, 397 in tomatoes, 387 in wheat, 395 citri, 60 in citrus, 384, 385 infectoria, 60, 62 in sorghum, 398 in wheat, 396

padwickiii, 137, 138 passiflorae, in passionfruit, 390 tenuis, 60, 62 tenuissima, 60, 62 Alternariol, 62 Alternariol monomethyl ether, 62 Altertoxin, 62 Anaerobic growth of fungi, 7 Anamorph, 15 Apples, spoilage of, 385, 386 Botrytis cinerea, 69 Epicoccum nigrum, 90 Penicillium brevicompactum, 233 Penicillium expansum, 245 Penicillium funiculosum, 264 Penicillium solitum, 258 Arthrinium, 58, 64, 132 phaeospermum, 64 state of Apiospora montagnei, 65 Ascomycete-conidial fungus connections, 15 Ascomycetes, 53 Ascomycotina, 13-15 Aspergillus, 277, 278, 279, 295 aculeatus, 277, 295 in onions, 391 alliaceus, 320 alutaceus, 317, 321 amstelodami, 282 awamori, 313 candidus, 277, 279, 297, 317 in cereals, 402 in cheese, 421 in flour and pasta, 403 heat resistance, 6 in maize, 402 in peanuts, 406 in rice, 403 in soybeans, 405 carbonarius, 277, 279, 299 in coffee beans, 410, 411 in dried vine fruit, 413

503

504 Aspergillus (cont.) in grapes, 388 ochratoxin A, 388, 411 chevalieri, 285 classification, 277 clavatus, 277, 279, 302 colony diameters, 278 fischeri, 293 fischerianus, 293 flavipes, 277, 279, 303 flavus, 277, 279, 305, 315, 322, 329 in Brazil nuts, 399, 409 in cashews, 399, 409 in coffee beans, 411 in copra, 400 in figs, 414 in maize, 397, 406 in peanuts, 398, 406, 407 in pistachios, 400, 409 in rice, 38, 396, 402, 403 in salt fish, 416, 417 in sorghum, 398 in soybeans, 397, 405 in spices, 410 in sunflower seed, 398 in tree nuts, 399, 401, 408, 409 fonsecaeus, 299 fumigatus, 51, 277, 279, 311 in maize products, 405 in rice, 402, 403 in tree nuts, 399, 408, 409 glaucus, 291 japonicus, 277, 279, 297 key to common species, 277 manginii, 291 nidulans, 279 niger, 277, 279, 313 in chickpeas, 406 in cocoa, 411, 412 in coffee beans, 410, 411 in dried vine fruit, 413 in figs, 389, 414 in grapes, 388 in maize, 404, 405 in onions, 391 in peanuts, 398, 406, 407 in pistachios, 400, 408, 409 in rice, 402, 403 in salt fish, 416, 417 in spices, 410 in tree nuts, 399, 407, 408 in yams, 392 niveus, 278, 279, 298, 315 nomius, 277, 279, 311 ochraceus, 278, 279, 317 in chickpeas, 406 in coffee beans, 410, 411 in dried fruit, 413 in figs, 414 in pistachios, 408 oryzae, 277, 279, 306, 310

Index parasiticus, 277, 279, 306, 321 in peanuts, 407 in tree nuts, 408 penicillioides, 278, 279, 323, 325 in cereals, 402 in confectionery, 415 in flour, 403 in fruit cakes, 414 in salt fish, 416 in processed meats, 417 pseudoglaucus, 287 pulchellus, 299 repens, 287 reptans, 287 restrictus, 278, 279, 324, 325 in cereals, 403 in fruit cakes, 414 in jams, 412 in maize, 404 in nuts, 409 in soybeans, 405 in rice, 403 ruber, 289 rubrobrunneus, 289 sejunctus, 289 sojae, 306, 322 steynii, 278, 279, 320 sydowii, 278, 279, 329 in cashews, 399, 409 in rice, 396 in salt fish, 416 in spices, 410 tamarii, 278, 279, 328 in Brazil nuts, 409 in cashews, 399, 409 in cocoa, 411, 412 in coffee beans, 411 in maize, 404 in peanuts, 406, 407 teleomorphs, 276 terreus, 278, 279, 330 tubingensis, 313, 314 ustus, 277, 279, 332 versicolor, 278, 279, 327, 333 vitis, 282 wentii, 278, 279, 336 westerdijkiae, 278, 279, 317, 320 in coffee beans, 411 ochratoxin, 411 Aspergillus flavus and parasiticus agar (AFPA), 28, 307, 319, 322, 329, 423 ATP, estimation of, 37 Aureobasidium, 57, 65 pullulans, 65 in meat, 394 Aurofusarin, 104, 117

B Baked goods, mycoflora of, 403-404 Eurotium chevalieri, 287

Index Balkan endemic nephropathy, 261 Bananas, spoilage of, 390 Alternaria alternata, 62 Colletotrichum gloeosporioides, 82 Fusarium oxysporum, 108 Fusarium semitectum, 113, 114 Fusarium subglutinans, 120 Fusarium verticillioides, 121, 122 Lasiodiplodia theobromae, 127 Nigrospora spherica, 133 Pestalotiopsis guepini, 134 Phoma sorghina, 135 Barley, mycoflora of, 395 Alternaria alternata, 62, 64 Arthrinium, 65 Aspergillus clavatus, 302 Chaetomium, 73 Cladosporium, 76, 79, 81 Fusarium culmorum, 100 Fusarium graminearum, 104 Fusarium sporotrichioides, 118 Geotrichum candidum, 124 Penicillium hordei, 249 Penicillium verrucosum, 261 Basidiomycotina, 15 Basipetospora, 340 halophila, 341, 350 in salt fish, 416 Beans, mycoflora of, 391, 397, 405 Rhizopus stolonifer, 160 Beauvericin, 102, 108, 110, 112, 117, 119 Beer, spoilage of, Brettanomyces bruxellensis, 358, 362 Berries, spoilage of, 389 Bettsia, 340, 343 alvei, 343 Biltong, mycoflora of, 412 Biomass, estimation of fungal, 34 ATP estimation, 37 chitin assay, 34 conductimetry, 37 ELISA, 39 ergosterol assay, 35 fluorescent antibodies, 40 immunological techniques, 38 impedimetry, 37 latex agglutination, 39 molecular methods, 40 PCR, 40 volatiles, 37 Biosystematics, 11-17 Bipolaris, 58, 67, 85 australiensis, 67 bicolor, 67 maydis, 67 oryzae, 67 setariae, 67 spicifera, 67 Botryodiplodia theobromae, 125

505 Botryosphaera rhodina, 125, 126 ribis, 386 Botrytis, 58, 68 allii, 69 in garlic, 391 byssoides, 69 cinerea, 68 in beans, 391 in berries, 389 in carrots, 392 in grapes, 388 in kiwifruit, 389 in peas, 391 in pome fruits, 385 in stone fruit, 386 state of Sclerotinia porri, 69 Bread, spoilage of, 403 Chrysonilia sitophila, 73, 74 Endomyces fibuliger, 88 Hyphopichia burtonii, 125 Moniliella suaveolens, 131 Penicillium roqueforti, 256 Pichia membranaefaciens, 371 Brettanomyces bruxellensis, 360, 361 intermedius, 360 Brines for olives, pickles and cheese, spoilage of, 358, 364, 365, 366, 371, 372 Butenolide, 102, 104 Butter, spoilage of, 394 Candida parapsilosis, 364 Penicillium thomii, 207 Rhodotorula mucilaginosa, 373 Byssochlamys, 169, 170 in cream cheeses, 393 fulva, 2, 5, 170, 171, 184 in heat processed acid foods, 418 heat resistance, 5, 171, 173 nivea, 6, 170, 173 oxygen requirements, 7, 171 spectabilis, 170, 184

C Cakes, spoilage of, 404 Eurotium rubrum, 291 Penicillium crustosum, 241 Xeromyces bisporus, 355 Calonectria, 90 Candida, 360, 361 chodatii, 124 famata, 364 holmii, 366 krusei, 360, 361, 362 mogii, 379 parapsilosis, 360, 361, 363 in cheese, 420 in pserved foods, 419 in yoghurt, 393

506 Candida (cont.) pelliculosa, 369 valida, 370 Capsicums, spoilage of, 387 Alternaria alternata, 62 Rhizopus stolonifer, 160 Carbon dioxide, tolerance of fungi to, 7 Carnation leaf agar (CLA), 43 Carpenteles, 175 Carrots, spoilage of, 392 Rhizopus stolonifer, 160 Cashews, mycoflora of, 399, 409 Aspergillus flavus, 309 Cassava, mycoflora of, 392, 393 Penicillium oxalicum, 218 Cephalosporium acremonium, 56 Cereals, mycoflora of Alternaria alternate, 60, 62 Aspergillus versicolor, 335 Chaetomium globosum, 73 Cladosporium, 78 Curvularia lunata, 84 Drechslera, 85 Endomyces fibuliger, 88 Fusarium avenaceum, 97 Fusarium culmorum, 100 Fusarium equiseti, 102 Fusarium graminearum, 104 Fusarium oxysporum, 108 Fusarium poae, 110 Fusarium sporotrichioides, 117, 118 Fusarium spp, 91, 92 Nigrospora oryzae, 132 Penicillium aurantiogriseum, 231 Penicillium chrysogenum, 237 Penicillium verrucosum, 261 Pestalotiopsis, 133 Phoma, 135, 136 Trichothecium roseum, 142 Cereals, spoilage of, 395-397, 402-406 Chaetoglobosin, 73 Chaetomium, 57, 70 brasiliense, 70 funicola, 70, 71 globosum, 70, 72 in cashews, 399, 409 in rice, 396 in soybeans, 397 in spices, 410 Cheese, manufacture, 419-421 Debaryomyces hansenii, 366 Penicillium camemberti, 235 Penicillium commune, 238 Penicillium roqueforti, 256 Cheese, mycoflora and spoilage of, 419-421 Aspergillus versicolor, 335 Candida parapsilosis, 364 Cladosporium, 75, 77, 78, 81 Debaryomyces hansenii, 366 Eurotium amstelodami, 284 Eurotium herbariorum, 292

Index Eurotium repens, 289 Fusarium oxysporum, 108 Geotrichum candidum, 124 Moniliella suaveolans, 131 Penicillium commune, 238 Penicillium crustosum, 241 Penicillium glabrum, 200 Penicillium roqueforti, 256 Pichia membranaefaciens, 371 Rhodotorula mucilaginosa, 373 Chickpeas, mycoflora of, 398, 405, 406 Chitin assay, 34-35 Chloroanisole formation in foods, 233, 411 Aspergillus versicolor, 336 Eurotium repens, 289 Paecilomyces variotii, 185 Penicillium chrysogenum, 236 Chocolates, spoilage of, 415 Chrysosporium inops, 347 Chrysosporium xerophilum, 347 Xeromyces bisporus, 355 Zygosaccharomyces rouxii, 382 Chrysonilia, 57, 73 sitophila, 51, 73, 74 Chrysosporium, 340, 342 farinicola, 343 in chocolate, 415 in coconut, 409 fastidium, 9, 344, 346 inops, 343, 345 in confectionery, 415 isolation techniques, 30 key to xerophilic species, 343 in prunes, 413 xerophilum, 343, 347 in coconut, 409 Citreoviridin, 197 from Eupenicillium ochrosalmoneum, 182 from Penicillium citreonigrum, 197 Citrinin, 129 from Monascus ruber, 129 from Penicillium citrinum, 208 from Penicillium verrucosum, 261 Citrus, spoilage of, 384-385 Alternaria, 61, 62 Fusarium oxysporum, 108 Geotrichum candidum, 123 Penicillium digitatum, 242 Penicillium italicum, 251 Penicillium ulaiense, 251 Cladosporium, 57, 75 butyri, 131 chodati, 124 cladosporioides, 75, 78, 80, 81 in cheese, 419 in chilled meat, 394 in coconut, 409 in low salt margarine, 394 in rice, 396 in salt fish, 416 in soybeans, 397, 405

Index in tree nuts, 399, 407 in wheat, 395, 396 in grapes, 389 herbarum, 75, 76, 77, 78 in cheese, 419, 420 in chickpeas, 398 in margarine, 394 in meat, 394 in tomatoes, 387 in tree nuts, 408 key to included species, 75 macrocarpum, 75, 78 sphaerospermum, 75, 76, 80 in citrus, 385 in maize products, 405 in strawberries, 389 suaveolens, 131, 394 Classification of fungi, 11, 16-17 Claviceps purpurea, 1 Cochliobolus lunatus, 83 Cocoa beans, mycoflora of, 411-412 Aspergillus fumigatus, 312 Candida krusei, 363 Kloeckera apiculata, 368 Coconut cream agar, 28 Coconut, spoilage of, 400, 409 Aspergillus niger, 315 Chrysosporium farinicola, 344 Chrysosporium xerophilum, 347 Coelomycetes, 54 Coffee beans, mycoflora of, 410-411 Aspergillus ochraceus and related species, 320 Ochratoxin in, 410, 411 Coffee, off flavours, 411 Aspergillus versicolor, 336 Colletotrichum, 57, 81 acutatum, 81, 82 in berries, 389 circans, 81 coccodes, 81 dermatium, 82 gloeosporioides, 81 in avocados and mangoes, 390 higginsianum, 81 lindemuthianum, 81 musae, 81, 82 in bananas, 390 Colony characters, 45 diameters, 45 Commodities, tropical, mycoflora of, 396-398 Chaetomium brasiliense, 70 Chaetomium funicola, 72 Cladosporium cladosporioides, 77 Concentrated foods, spoilage of, 412-416 Conductimetry, 37 Confectionery, spoilage of, 415 Chrysosporium inops, 347 Xeromyces bisporus, 355 Zygosaccharomyces rouxii, 382 Consistency, effect on growth, 8

507 Copra, mycoflora of, 400 Cream cheese, spoilage of, 393, 419, 421 Creatine sucrose agar (CREA), 42, 224 Creatine sucrose neutral agar (CSN), 224, 423 Cucumbers, spoilage of, 388 Penicillium oxalicum, 218 Culmorin, 100, 104 Culture mites, 50 Cultures, examination, 45 Cunninghamella, 148, 149 berttholletiae, 149 echinulata, 149 elegans, 149 Curvularia, 58, 67, 82 lunata, 83 lunata var. aeria, 83 pallescens, 83, 8

【#9】Cách Chăm Sóc Hoa Thủy Tiên Nước

Thân dài tới 120 m buông rũ xuống. Tách cây ngâm chậu lan vào nước 15 ph cho rễ mềm gỡ cây ra khỏi chậu dùng dao thật sắc tách từng 3 giả hành thành 1 đơn vị với những cây quí người ta 1 giả hành nếu ta chưa có nhiếu kn thì không nên.

Hoa Thủy Tiên Hoa đẹp Rực Rỡ Cho Ngày Tết Nhiều May Mắn

Lá mọc đối cách dài 8 12 cm rộng từ 4 7 cm.

Cách chăm sóc hoa thủy tiên nước. Cách trồng mai vàng chuẩn bị ghép mai vàng có nhiều loại có loại hoa to loại hoa nhỏ nhưng đa số có 5 cánh và đều mau tàn hoa sẽ rụng sau 1 ngàydo vậy những ngày tết chúng sẽ rụng nhiều nhất là khi chúng ta để nó trong nhà. Nếu bạn muốn chơi tết bằng loại hoa có nét đẹp rực rỡ kiêu hãnh và duyên dáng thì hoa trà my là lựa chọn tuyệt vời. Hoa to tới 10 cm mọc từ 1 3 chiếc ở các đốt đã rụng lá nở vào mùa xuân.

Cách tách lan chiết lan và chăm sóc lan. Là cơ hội đầu tư sinh lời cao. Bệnh viện stbernard với đội ngũ y bác sĩ ưu tú giàu kinh nghiệm cùng với trang thiết bị hiện đại là địa chỉ uy tín chăm sóc sức khỏe gia đình bạn.

Hoa tóc tiên hoa đẹp. Cá cảnh thái hòa mua bán cá cảnh cá rồng cá đĩa cá koi cá thủy sinh và các loại bể cá cảnh bể cá rồng bể thủy sinh bể treo tường bể trung quốc. Phương cách nuôi dưỡng chú cá rồng chăm sóc cá rồng cách chăm sóc cá rồng tốt kỹ thuật nuôi cá rồng.

Hoa trà my hoa tết độc đáo cho mọi nhà. Cây cảnh việt nam là website đầu tiên tại việt nam cung cấp thông tin và tư vấn kỹ thuật cây cảnh nghệ thuật bonsai. Dự án nghĩa trang sala garden mô hình hoa viên sinh thái đầu tiên tại việt nam.

Hoa tóc tiên huệ mưa rain lily không kiêu sa nhưng rực rỡ nhỏ bé khiêm nhường nhưng hiên ngang trước mưa gió bão bùng.

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【#10】Cách Trồng Và Chăm Sóc Mai Vàng Đúng Kỹ Thuật Nhất

Mai là cây hoa kiểng dễ sống, sống mạnh và được coi là giống cây dễ trồng nhất. Nó không quá kén đất trồng. Bằng chứng là trên các loại đất khác nhau như đất thịt, đất cát pha, sét pha, đất phù sa, đất đỏ bazan, thậm chí đất có lẫn đá sỏi…mai vẫn phát triển tốt. Tuy nhiên, mai kỵ đất bị úng thuỷ, thường xuyên ngập lụt. Rễ mai dài bởi vậy nước ngập lâu dài sẽ khiến cây bị héo úa và chết dần. Trong rễ mai, rễ bàng (mọc quanh đoạn cổ rễ) có sức sống mạnh nhất. Bị chặt đứt, chúng lại mọc ra. Vì vậy, bộ rễ bàng đóng vai trò quan trọng với việc sinh trưởng và phát triển của cây mai.Nhân giống hữu tính: Đây là cách trồng mai bằng hạt. Cách trồng này có ưu điểm là số lượng mai con nhiều, không tốn kém, mất ít công sức. Tuy nhiên, cây mai thường không mang những đặc tính tốt của cây mẹ (hoa nhỏ, ít cành hơn, màu sắc có khi khác với cây mẹ…).

Điều quan trọng nhất với mai kiểng là dáng cây và việc ra hoa đúng kỳ.Cành lá quá tốt sẽ gây ra ức chế với việc trổ hoa. Cây còi cọc quá thì hoa ít, không đẹp. Mai kiểng được trồng trong chậu với lượng đất giới hạn, nên việc bón phân, chăm sóc cho mai hết sức cần thiết.

Người trồng mai kiểng còn chú ý đến môi trường sống của mai. Mai là loại cây thích hợp với khí hậu nóng ấm, từ 25o – 30o, có thể chịu được nhiệt độ cao trong nhiều ngày, nhiều tháng. Tuy nhiên, ở những nơi nhiệt độ thấp (dưới 10o), mai sinh trưởng kém.

Kỹ thuật trồng và chăm sóc mai

Mỗi giống cây đều có một cách trồng riêng. Có giống đòi hỏi phải trồng với kỹ thuật cao, (trồng đúng kỹ thuật cây mới cho năng suất cao). Nhưng nhiều loại cây có cách trồng giản dị. Mai là một loại cây như vậy. Tuy nhiên, đó là cách trồng để cây mai sống và ra hoa. Còn trồng theo cách ghép cành, uốn thế để có cây mai kiểng cổ, cây mai ghép nhiều màu, hoặc cây mai bonsai tuyệt đẹp thì lại là một việc khác. Nó đòi hỏi người trồng phải có kỹ thuật, hiểu biết về loài cây này.

1. Lên luống và mương rãnh thoát nước

Cây mai không hợp với những vùng đất thấp, đất có mạch nước ngầm dâng quá cao, đất thường xuyên bị ngập úng vào mùa mưa. Nếu trồng mai ở thế đất như trên, cần lên luống. Thông thường, bề ngang luống rộng từ 1-1,2m. Luống này sẽ dùng để ương mai con, khi lướn bứng trồng vào chậu. Giữa hai luống mai sát nhau nên có mương, rãnh thoát nước, tránh ngập úng cho vườn mai.

2. Nhân giống

a).

b). Nhân giống vô tính: Đây là cách trồng mai được tiến hành bằng việc chiết cành, ghép cành, hoặc giâm cành. Cách nhân giống này giúp cây con giữ được trọn vẹn những đặc tính của cây mẹ. Tuy vậy, với cách nhân giống này, mai không thể sản xuất đại trà với số lượng lớn.

* Chiết cành: Chọn một cành nhỏ của cây mai mẹ, cắt một khoanh vỏ có chiều dài khoảng 3-4 phân, cố tránh đừng để vết cắt phạm vào phần gỗ bên trong, bóc khoanh vỏ đó đi. Sau đó, dùng hỗn hợp đất với phân chuồng hoai nhào dại cho dẻo rồi ốp chặt vào xung quanh vết cắt, bên ngoài dùng vải dày hay bao bố hoặc xơ dừa bó lại cho thật chặt. Hàng ngày phải năng tưới nước cho bầu đất đó được ẩm cho đến vài ba tháng sau, khi bầu đất có nhiều rễ con bắn ra ngoài là lúc cắt nhánh đó rời khỏi cây mẹ.

* Ghép cành (tháp cành, tháp cây): là dùng cành của cây mẹ đem ghép vào cây mai khác để tạo cây mai mới mang những đặc tính của cây mai mẹ.

Có một cách ghép khác là ghép mắt, là lấy mắt lá, chồi non từ cây mẹ để ghép sang một cây khác làm gốc ghép.

* Ghép tam giác: Lấy một cây mai làm gốc ghép, lựa một chỗ trên gốc cây để ghép cành hay ghép mắt, dùng mũi dao nhọn rạch một hình tam giác nhỏ tương đương hột bắp rồi bóc lớp vỏ đó ra.

Dùng dao bén tách ra một chồi nhỏ hay một mắt lá của cây mai mẹ đem áp vào chổ tam giác vừa được lột vỏ của gốc ghép. Sau đó, dùng dây vải hoặc băng keo băng mắt ghép lại. Sau vài tuần, thấy chồi ghép hay mắt ghép xanh tươi có nghĩa là thành công.

Một gốc ghép có thể ghép được nhiều chồi hay nhiều mắt ghép. Ta thấy một cây mai ghép có nhiều màu hoa khác nhau chính là do cách ghép này.

* Ghép nêm: Dùng dao vạt hình cái nêm trên cành ghép và hình lỗ nên trên gốc ghép (hay làm ngược lại) rồi ráp khít hai bộ phận trên lại với nhau. Yêu cầu là cành ghép và gốc ghép phải có đường kính bằng nhau hay gần bằng nhau và cả 2 cây phải có độ tuổi ngang nhau mới tốt.

Đặt 2 mối khít với nhau, ta dùng dây cao su hoặc dây nylon quấn chặt bên ngoài vết ghép cho chắc chắn.

Nên ghép cây vào mùa mưa, vì đây là mùa cây đang dồi dào sinh lực. Tại gốc ghép, chọn nơi vỏ cây tươi tốt để tạo chỗ ghép, như vậy mắt ghép mới hy vọng đạt được thành công, vì nơi ấy nhựa nguyên lưu thông tốt. Việc ghép phải thực hiện càng nhanh càng tốt, để lâu nhựa sẽ khô, ghép không có kết quả.

3. Chăm sóc mai

* Tưới nước: Cây mai tuy chịu nắng hạn, nhưng không có nghĩa là có khả năng chịu hạn cao. Trong mùa nắng, ta nên chăm lo tưới nước. Với mai trồng đại trà ngoài vườn, mỗi ngày hoặc cách ngày tưới nước một lần mới tốt. Tưới thẳng vào gốc và xịt nước với tia nhỏ lên khắp tán lá lại tốt hơn. Nên tưới vào lúc sáng sớm (trước 9 giờ) hoặc tưới vào lúc chiều mát.

Vào mùa mưa, mai trồng trong vườn khỏi tưới cũng được, trừ trường hợp nhiều ngày nắng gắt kéo dài thì phải tưới nước để giữ đất đủ ẩm. Mai kiểng trồng trong chậu thường bị khô nước vì đất chứa trong chậu quá ít nên không giữ ẩm được lâu. Do đó, mai kiểng trồng trong chậu phải tưới nước mỗi ngày, ngày tưới 2 lần (sáng, chiều).

Phải chú ý đến độ rút nước của từng chậu, nếu thấy có tình trạng úng nước phải dùng que nhỏ thông ngay, nếu để lâu cây mai sẽ bị chết vì bộ rễ bị hư.

* Bón phân: Trồng mai phải bón phân, nhất là đối với cây trồng trong chậu, sau khi tỉa cành tạo dáng cho chúng ta cần bón phân cho mai sinh trưởng tốt về cành lá.

Lúc này yêu cầu đạm và lân nhiều hơn, kali ít cũng được. Có thể dùng phân Đầu Trâu NPK 20-20-15TE, xới đất lên bón, lấp đất lại. Lượng bón mỗi lần không cần nhiều: khoảng 40-50 g/chậu chứa 50-60kg đất (đối với cây trồng ngoài đất lượng bón tương tự như trong chậu nhưng bón xa gốc cây, khoảng rìa ngoài của tán cây), tưới đủ nước thường xuyên (trong mùa khô). Mỗi tháng bón 2-3 lần, quan sát thấy cây ra lá, cành lá xum xuê là được. Nếu thấy lá quá đậm thì giảm số lượng cũng như số lần bón xuống.

Vào mùa mưa từ tháng 6-10 dương lịch, dùng NPK Đầu Trâu 13-13-13TE để bón, mỗi lần bón 40-50g/chậu chứa 50-60kg đất, 15-20 ngày bón một lần. Bón các loại phân trên đã cung cấp đầy đủ các chất đa lượng và vi lượng cho mai. Tuy nhiên khi thay đất hoặc sau 3-4 tháng kể từ khi thay đất có thể bón thêm phân chuồng: phân bò, heo, gà vịt đã ủ kỹ kết hợp với tro trấu cũng rất tốt.

Khi kết thúc mùa mưa: khoảng giữa tháng 11 dương lịch. Tiến hành xem lại dáng cây, cành lá đã thỏa mãn chưa. Có thể tỉa lại một lần nữa rồi chỉ tưới nước dưỡng cây.

* Diệt cỏ dại, bắt sâu: Cỏ dại tranh ăn chất bổ của phân được bón vào đất, vì vậy cần phải tiêu diệt ngay. Nên diệt cỏ vào trước mùa mưa hàng năm. Về sâu bệnh, cây mai có đặc tính kháng bệnh cao, nên ít bị sâu rầy phá hại, thế nhưng không phải là không có. Chúng ta nên quan sát, nếu phát hiện có sâu rầy thì nên tận diệt ngay. Một số sâu, rầy chính hại cây mai: Sâu đục thân, rầy bông, sâu tơ, sâu nái.

4. Trẩy (lặt) lá mai

Đây là việc làm giúp mai nở hoa đúng vào dịp Tết. Thời gian để trẩy lá mai không nhiều, tiến hành xong trong ngày mới tốt. Nếu kéo dài thì mai sẽ nở hoa không đúng ngày. Muốn cây mai trổ hoa nhiều, người trồng mai cần trẩy hết là non, lá già, chú ý không làm gẫy ngọn cành.

Có 2 cách trẩy lá mai: Cách thứ nhất, ta cầm lá trẩy ngược ra sau. Cách này tốn ít sức, nhưng dễ kéo theo một đoạn dài vỏ, dễ làm hư hại đến cành hoa và nụ hoa. Cách thứ hai, ta cầm lá kéo theo chiều của chiếc lá. Cách này không làm xước vỏ, nhưng tốn nhiều công sức. Hơn nữa, đối với những đọt non, cách này dễ gây ra đứt đọn do kéo quá sứ

Kỹ thuật trồng mai vàng

Tên khoa học: Ochna integerrina

Từ ngàn xưa, người dân Nam bộ đã xem hoa mai là biểu tượng của xuân về, hình ảnh hoa mai vàng khoa sắc, đâm chồi nảy lộc với hàm ý mong một năm mới nhiều tài lộc và thịnh vượng. Vì lẽ đó hoa mai vàng luôn được ưa chuộng hơn hoa mai trắng. Tùy vào sở thích và mục đích sử dụng cây mai mà có những cách trồng mai khác nhau, có cách đòi hỏi phải trồng với kỹ thuật cao (trồng theo cách ghép cành, uốn để có cây mai kiểng cổ, cây mai ghép nhiều màu, hoặc cây mai bonsai) hay chỉ trồng giản dị trong đất để mai sống và ra hoa.

Cây mai có thể nhân giống bằng phương pháp hữu tính (trồng bằng hạt, thường mất từ 5 – 6 năm mới có thể sử dụng) hay phương pháp vô tính (chiết cành, ghép cành hoặc giâm cành, có thể sử dụng sau 2 – 3 năm).

Mai vàng trồng được quanh năm nhưng tốt nhất là gieo hạt vào tháng 2 âm lịch. Cây mai trồng vào chậu nên chọn cuối tháng 10 âm lịch của năm trước đến tháng 2 âm lịch của năm sau, chính là điều kiện tốt để hình mô sẹo và mọc chồi.

Hơn nữa, ánh sáng là yếu tố quan trọng trong quá trình sinh trưởng và phát triển của mai vàng, nên đảm bảo thời gian nhận ánh sáng được 6 tiếng trở lên. Những nơi có thời gian chiếu sáng quá ít, cây mai thường sinh trưởng kém và ra hoa ít.

Mai vàng thích hợp khí hậu nóng ẩm hoặc có thể chịu đựng ở nhiệt độ cao hơn trong nhiều ngày và nhiều tháng. Tuy nhiên, với những vùng có khí hậu mát lạnh dưới 10 0C thì mai sinh trưởng kém. Năm nào thời tiết cuối năm thay đổi như mưa nhiều hoặc giá lạnh thì cây mai cũng nở hoa không đúng ngày (nhiệt độ thích hợp nhất từ 25 0C – 30 0 C)

+ Gieo hạt: hạt chín (có màu đen) còn tươi thì tiến hành gieo ngay, có thể đạt tỉ lệ nảy mầm trên 95%. Cứ 1 m 2 gieo được 100 hạt, cây con có chiều cao 10 cm có thể bứng ra trồng trong chậu hoặc giỏ tre.

+ Trồng chậu: nếu chậu nhỏ có thể xếp 4 chậu/1m 2, chậu lớn thì xếp 1chậu/ 1 – 2 m 2 nhằm đảm bảo đủ ánh sáng cho cây.

Mai vàng không kén đất, có thể trồng trên các loại đất thịt, đất cát pha, sét pha, đất phù sa, đất đỏ bazan, hoặc đất có lẫn đá sỏi. Tuy nhiên, với vùng đất thấp cần lên líp rộng 1 – 1,2 m, có rãnh thoát nước để mai không bị úng ngập khi mưa hay nước ngầm dâng cao làm úng, thối rễ. Riêng mai trồng chậu nên bổ sung tro trấu, xơ dừa, vỏ đậu phộng,… và cát sẽ giúp thoát nước rất tốt.

Phân hữu cơ rất được ưa chuộng và được xem là loại phân chính như: phân chuồng, rơm rạ mục, mùn dừa, đầu tôm, đầu cá, xác đậu nành,… hoặc có thể bổ sung phân Dynamid và phân lân hữu cơ sông Gianh. Phân hữu cơ giúp cho mai phát triển bền vững, tạo nhiều nụ hoa và có tác dụng làm tăng độ pH trong chất trồng.

Kết hợp dùng phân tổng hợp NPK 30-10-10 cho cây vào đầu năm. Từ giữa năm đến tết thì bón vài lần phân NPK 20-20-15 để giúp mai kết nụ và nở bông tốt.

Có thể dùng thêm phân bón lá để giúp cây tăng trưởng nhanh, tạo nhiều nụ và hoa.

Lịch bón phân tham khảo cho mai trồng chậu

Chú ý: Đối với mai ghép trồng chậu bón phân từ tháng 2 ÂL đến 15/9 ÂL, một tháng bón phân một lần. Từ tháng 10 đến cuối tháng 11 âm lịch, không bón phân vào gốc và hạn chế tưới nước để chuẩn bị lặt lá.

Liều lượng sử dụng phân hữu cơ Dynamid từ 10gr đến 100gr/ chậu lớn (đường kính 80 – 100 cm). Phân hoá học NPK từ 10gr đến 50gr/ chậu lớn, Lân hữu cơ sông Gianh từ 10gr đến 30gr/ chậu lớn.

Lịch bón phân tham khảo cho mai trồng ngoài đất

Không nên bón quá nhiều loại phân cùng một lúc, cây dễ bị chết vì ngộ độc hoặc bội thực.

Chú ý: – Bón lót trước khi trồng bao gồm phân hữu cơ (phân chuồng hoai hoặc hữu cơ Dynamid) và bón thúc có 3 lần trong năm.

– Liều lượng sử dụng phân hữu cơ Dynamid từ 100 kg/1.000m 2, phân hoá học NPK từ 10 – 15 kg/1000 m 2, Lân hữu cơ sông Gianh từ 100kg/1.000m 2.

Các đối tượng thường xuyên gây hại trên cây mai có thể kể đến:

Có thể sử dụng một trong các loại thuốc trừ sâu như sau: Malvate 21EC; Trebon 10EC; Confidor 100SL; Admire 050EC; Regent 5SC, Vimite 10ND; Bifentox 30ND; Virigent 800WG…

b. Nhện đỏ (Tetranychus sp.)

Khi phát hiện có nhiều nhện trên cây có thể dùng một trong các loại thuốc: Danitol 10EC; Comite 73EC; Pegasus 500SG; Ortus 5SC; Cascade 5EC;

Nissuran 5EC; Sirbon 5EC; Kelthane 18,5EC…

c. Rệp sáp (Dysmiccocus sp)

Có thể dùng tay giết rệp hoặc khi cần thiết thì sử dụng một trong các loại thuốc: Pyrinex, Supracide, Polytrin, Monster…

d. Sâu ăn lá (Delias aglaia)

Có thể sử dụng thuốc trừ sâu như: SecSaigon 5EC hoặc 10EC; Diaphos 5EC; Sagothion 50EC; Padan 95SP; Fastac 5EC,…

đ. Bệnh mốc cam (do nấm Coniothyrium fuckelli)

Nên định kỳ tỉa cành, cắt bỏ các cành bị gẫy hoặc bị bệnh. Sau khi tỉa cành phun thuốc Daconil, Zineb, hoặc thuốc gốc đồng,…

e. Bệnh gỉ sắt (do nấm Phragmidium mucronatum)

Tỉa bỏ các cành lá bệnh rồi tập trung tiêu huỷ. Bón lân và kali tăng sức chống bệnh cho cây. Chỉ tưới nước vừa phải. Phun thuốc Bayfidan, Score, Anvil, Zineb, Carbendazim,…

f. Bệnh cháy lá (do nấm Pestalotia funereal)

Bón phân đầy đủ, cân đối tỉ lệ N-P-K, ngắt bỏ lá già, lá bệnh, định kỳ phun thuốc gốc đồng và phân bón lá cho cây.

g. Bệnh vàng lá do tác nhân bệnh sinh lý

Bón đầy đủ phân khi có hiện tượng vàng lá, nên kết hợp phun xịt phân bón lá có chất vi lượng, cây sẽ mau hết bệnh.

h. Bệnh đốm lá (do nấm Pestalotia palmarum)

Dùng biện pháp phòng trừ tổng hợp như: mật độ trồng thích hợp để cây mai được thông thoáng, vệ sinh vườn bằng cách cắt tỉa thu gom lá bị bệnh tiêu huỷ để tránh lây lan, bón phân cân đối, cần tăng cường bón thêm phân hữu cơ và kali cao giúp cây kháng bệnh. Khi cây bệnh có thể dùng thuốc hoá học: Viben C BTN, phun ướt đều cả hai mặt lá, cần lập lại 2-3 lần, sau 5-7 ngày để trị bệnh. Hoặc có thể phun từ 10-15 ngày/ lần để phòng trừ bệnh.

i. Bệnh đốm đồng tiền do tác nhân địa y

Không nên trồng hoặc sắp xếp những chậu mai trong vườn quá dày, quá gần nhau, dưới tán lá và gốc cây cần nhận đủ ánh sáng mặt trời. Cần để vườn mai thông thoáng, khô ráo. Thiết kế mặt liếp để trồng mai (hoặc để đặt chậu mai) theo hình mai rùa, xẻ rãnh thoát nước để nước không đọng lại trên mặt vườn trong mùa mưa để vườn luôn được khô ráo.

Đối với những gốc mai đã bị đốm bệnh xuất hiện nhiều, dày đặc, có thể dùng bàn chải cọ rửa sạch những đốm bệnh trên thân, cành.

Dùng nước vôi hoặc dung dịch thuốc Bordeaux 1% quét lên thân cây vào đầu màu mưa, cũng có tác dụng ngăn chặn bệnh xâm nhập và lây lan. Ngoài ra bạn có thể dùng một số loại thuốc gốc đồng như Copper – B, Coc 85; Copper – Zinc hoặc Zinccopper… xịt ngừa lên chỗ thường hay bị bệnh trên thân, cành.

* Đầu vụ cần làm tốt để không ảnh hưởng cả một vụ mai như dọn cỏ, bón phân, thay đất, tỉa cành, tạo dáng cây cần có tay nghề cao mới làm được.

* Cuối vụ lặt lá mai đúng lúc là khâu rất quan trọng. Cần dựa vào tình trạng sinh trưởng, đất trồng, nước tưới, độ tuổi của cây, điều kiện thời tiết khí hậu, mai trồng trong chậu hay trong vườn…

Những năm thời tiết không thay đổi, thường lặt lá mai vào rằm tháng 12 âm lịch. Nhưng nếu thời tiết có sự thay đổi thì tùy theo mức độ nắng nóng hoặc lạnh có thể tiến hành lặt lá mai như sau:

– Có nắng nóng nhiều hoặc có gió chướng mạnh thì mai sẽ nở sớm hơn vì thế nên lặt lá mai trễ hơn. Tùy theo kích thước của nụ mai mà có thể lặt lá mai vào khoảng từ ngày 17-20 tháng chạp.

– Năm mưa nhiều và chấm dứt muộn, thời tiết tháng chạp lạnh nhiều, ít gió chướng thì mai sẽ nở trễ hơn, vì thế nên lặt lá mai vào trước ngày rằm tháng chạp, khoảng từ ngày 10-13 tháng chạp tùy theo kích thước nụ mai đã lớn hay nhỏ.

– Năm có tháng nhuận mai cũng nở sớm hơn vì thế nên lặt lá mai trễ hơn.

– Khi trời lập xuân sớm thì mai sẽ nở sớm, năm đó nên lặt lá mai trễ hơn, ngược lại năm lập xuân trễ thì mai sẽ nở muộn hơn, năm đó nên lặt lá sớm hơn.

* Sau tết nguyên đán : nên chăm sóc ngay để cây phục hồi nhanh.

Nếu có đất vườn thì chuyển ngay mai từ chậu ra trồng trong đất.

Nếu không có đất vườn thì nên thay đất mới. Nên bỏ bớt khoảng 1/3 đất cũ trong chậu, thay bằng hỗn hợp 3 phần chất trồng mới, 1 phần phân hữu cơ. Hòa 15 – 25 gam phân NPK 20-20-15 trong 10 lít nước tưới đều vào gốc mai. Tiếp tục bón thúc và tưới nước định kỳ để mai phát triển tốt. Việc bón phân, thay đất là rất quan trọng vì nó cung cấp dinh dưỡng cho cây mai phát triển trong suốt một năm. Sau giai đoạn này, người trồng mai nên tiếp tục chế độ chăm sóc trong cả năm, nhất là việc thường xuyên tưới nước.

Trong mùa nắng nên tưới mỗi ngày để đất đủ ẩm, mùa mưa cần đảm bảo tiêu thoát nước tốt và chỉ tưới khi đất khô. Có thể tưới tràn, tưới phun lên cả thân lá hay tưới rãnh, tưới nhỏ giọt. Mai trồng trong chậu bốc thoát hơi nước nhanh nên cần tưới nhiều lần hơn so với mai trồng trong đất. Chỉ nên tưới vào sáng sớm hay chiều mát (khi trời không quá nắng). Không nên tưới quá đẫm vào buổi chiều tối vì rất dễ phát sinh sâu bệnh do ẩm độ ban đêm rất cao.

Kỹ thuật trồng, chăm sóc mai vàng để hoa nở đẹp, đúng tết

Cả tuần nay ngày nào cô em gái cũng bận rộn, săm soi mấy chậu mai, lặt lá cây nào trước, rồi tưới nước. Muốn có hoa mai nở kịp Tết người trồng hoa rất vất vả, bỏ bao nhiêu công sức tưới tắm, bón phân trong cả năm…, vậy mà nhiều khi kỹ thuật cũng bó tay trước thời tiết đỏng đảnh.

Nhân tìm đọc một số bài viết và tài liệu về chăm sóc mai vàng cho em gái, xin chia sẻ một bài viết hay trên báo Nông Thôn Ngày Nay và các trang mạng khác:

Chuẩn bị đất: Với những vùng đất thấp cần lên líp rộng 1-1,2m, có rãnh thoát nước để mai không bị úng ngập khi mưa hay nước ngầm dâng cao làm thối rễ mai. Xới đất cho tơi xốp, nhặt hết cỏ dại và gạch đá.

Bón lót: Bón phân bò, tro trấu với lượng 3-5kg hay phân hữu cơ Đầu Trâu với lượng 0,3- 0,5 kg cho mỗi hố trồng. Nếu trồng mai trong chậu cần trộn đất với phân theo tỷ lệ 3-4 phần đất, 1 phần phân hữu cơ. Rải một phần phân hữu cơ xuống hố, đặt cây, rải tiếp phân hữu cơ quanh gốc rồi lấp đất, lèn chặt.

Tưới nước: Mùa nắng nên tưới hàng ngày để đất đủ ẩm, mùa mưa cần đảm bảo tiêu thoát nước tốt và chỉ tưới khi đất khô. Mai trồng trong chậu bốc thoát hơi nước nhanh nên cần tưới nhiều lần hơn so với mai trồng trong đất. Chỉ tưới vào sáng sớm hay chiều mát, không nên tưới quá đẫm vào chiều tối vì dễ phát sinh sâu bệnh do độ ẩm quá cao vào ban đêm.

Bón phân thúc: Sau trồng 15-20 ngày, rễ mai đã ăn ra lớp đất mới, cần tưới phân thúc bằng cách hòa 15-25gam phân NPK 20-20-15+TE Đầu Trâu/10 lít nước tưới vào gốc nhằm thúc cho bộ rễ mai phát triển mạnh ngay từ đầu. Bón thúc bằng cách rải phân NPK 20-20- 15+TE hoặc NPK 13-13-13 Đầu Trâu quanh gốc với lượng 20-30 gam/cây, định kỳ 25-30 ngày/lần. Kết hợp xới đất để vùi lấp phân nhằm giảm bớt thất thoát phân do bay hơi, rửa trôi. Phun phân bón lá Đầu Trâu 501 định kỳ 7-10 ngày/lần. Sau 3-4 tháng từ khi trồng, bón 0,5-0,1 kg phân hữu cơ/cây. Cuối tháng 10 âm lịch cần giảm lượng phân và nước tưới để hạn chế tăng trưởng thân lá, chuẩn bị cho giai đoạn phân hóa mầm hoa. Phun phân bón lá Đầu Trâu 701 định kỳ 7-10 ngày/lần nhằm kích thích mai phân hóa mầm hoa tốt.

Biện pháp xử lý để mai vàng ra hoa đúng tết: Cần áp dụng đồng bộ: Bón phân – Xiết nước – Tuốt lá. Từ đầu tháng 10 âm lịch hạn chế bón các loại phân có hàm lượng đạm (N) cao. Từ giữa đến cuối tháng 11 âm lịch, dừng bón phân vào gốc và hạn chế tưới nước để chuẩn bị tuốt lá. Từ 7-10 tháng Chạp, nếu thấy mai sung sức, đã có nụ lớn thời tiết dự báo nắng ấm thì mai sẽ nở sớm, do vậy đối với mai 5 cánh cần tuốt lá vào khoảng 18-20 tháng Chạp. Ngược lại nếu cây mai không sung sức, mới xuất hiện nụ nhỏ, dự báo rét kéo dài thì phải tuốt lá khoảng ngày 13-16 tháng Chạp. Đối với mai nhiều cánh cần tuốt lá sớm hơn so với mai 5 cánh từ 4-6 ngày. Trước khi tuốt lá cần ngừng tưới nước 2- 3 ngày để lá bắt đầu đanh lại, gân lá nổi lên thì tuốt lá, đồng thời tưới lại thật đẫm và phun phân bón lá Đầu Trâu 701. Đúng “tết ông Táo”, nếu thấy hoa cái bung vỏ lụa là chắc chắn hoa nở đúng tết; Nếu hoa cái chưa bung vỏ lụa là mai nở muộn nên cần xiết nước (ngưng tưới), đem phơi ngoài nắng (nếu trồng chậu) sau vài ngày thì tưới thật đẫm trở lại bằng nước ấm (45-50 độ C) đồng thời phun phân bón lá Đầu Trâu 901 để kích thích mai nở sớm cho đúng tết. Nếu hoa cái đã bung vỏ lụa trước “tết Ông Táo” thì mai sẽ nở trước tết nên cần phải hòa 10-20 gam phân urea/10 lít nước để tưới. Đồng thời cần tưới bằng nước lạnh (có thể cho một ít nước đá vào) và dùng lưới bạt che nắng để hãm mai nhằm giúp hoa nở đúng tết. Đối với những năm nhuận, thường mai sẽ nở sớm hơn nên cần kéo dài thời gian bón phân thúc và tưới nước so với những năm thường để thời gian tăng trưởng thân lá lâu hơn, giúp mai nở đúng tết. Việc tuốt lá, phun phân bón lá cũng theo nguyên tắc trên. Từ cuối tháng 11, nếu có mưa bất thường thì mai sẽ nở sớm do đó cần chủ động nắm bắt dự báo để có thể làm dàn che hay phủ nilon che gốc để tránh mưa.

Chưng mai trong những ngày tết: Chậu mai phải để nơi thoáng mát, đủ sáng không nên để gần quạt hay chỗ có gió lùa vì sẽ làm mai mất nước nhiều rụng hoa và cả nụ sớm. Không nên để mai chỗ quá tối vì sẽ không đủ ánh sáng cho mai quang hợp, chồi sẽ vươn dài, lá ra nhanh, hoa rụng sớm. Nên tránh để mai gần bóng đèn có công suất lớn vì sẽ thừa sáng, nhiệt độ lại cao cũng làm mai nở nhanh, chóng tàn. Nếu là cành mai cắm trong bình cần phải thui gốc ngay sau khi cắt để giữ nhựa và hạn chế vi khuẩn gây thối cành. Thay nước sạch nhiều lần hoặc cho mỗi lít nước 1 viên Aspirin nhằm hạn chế vi khuẩn gây thối cành, tàn hoa.

Chăm sóc mai sau tết: Sau tết, mai rất mất sức nên cần chuyển mai từ chậu ra trồng trong đất. Nếu vẫn trồng trong chậu cần phải thay đất mới bằng cách bỏ bớt khoảng 1/3 đất cũ trong chậu, thay bằng hỗn hợp 3 phần đất mới, 1 phần phân hữu cơ. Hòa 15- 25 gam phân NPK 20-20-15 Đầu Trâu trong 10 lít nước, tưới đều vào gốc mai. Tiếp tục bón thúc và tưới nước, phun phân bón lá theo chu kỳ mới như đã nêu ở trên .

Sau Tết, mai bắt đầu tàn tạ và cần được chăm sóc, việc làm này sẽ tạo nền tảng cho cây ra hoa cuối năm sau, phát triển tốt và tránh sâu bệnh.

Độc giả Hoàng Minh gửi đến VnExpss bài viết về cách chăm sóc cây mai sau Tết, độc giả này khuyên mọi người nên chăm sóc cây từ trước rằm tháng riêng.

“Việc chăm sóc mai sau Tết chia là 3 loại cây: Cây trồng chậu chưng trong nhà, cây trồng chậu chưng ngoài sân và cây trồng đất.

Cây mai vàng. Ảnh: Caycanhviet.

Với cây trồng chậu chưng trong nhà:

Mai chưng trong mấy ngày Tết thường từ 28 đến mồng 6 Tết, cây không được tiếp xúc trực tiếp với ánh sáng mặt trời nên hiện tượng quang hợp không thực hiện bao nhiêu, lá phát triển mới thường có màu xanh nhợt nhạt, mỏng, cành phát triển dài và yếu. Chủ nhà nhiều khi không tưới một ít nước mỗi ngày mà có khi đổ cả bia, nước ngọt vào gốc mai.

Chưa kể đến việc dùng thuốc kích thích ra hoa, giữ hoa không rụng làm xáo trộn sinh lý của cây. Cây mai dồn nhựa ra để nuôi hoa đẹp, lại phải thiếu điều kiện sống trong một tuần lễ nên nhiều cây bị kiệt sức rất nhiều, nếu không chăm sóc tốt thì năm sau mai sẽ không còn hoa nữa.

Mai được đem ra ngoài càng sớm càng tốt, phải đặt cây nơi có bóng râm để lá không bị cháy khi tiếp xúc trực tiếp với ánh sáng mặt trời. Lặt bỏ tất cả các hoa dù nở hay chưa nở (cả nụ còn lại) để cây không mất dinh dưỡng nuôi đài hoa tạo hạt, một số lá tạo trong lúc nở hoa dù xấu cũng để nguyên như vậy khoảng hơn một tuần cho cây hồi phục.

Với cây mai chưng ngoài sân hoặc cây mai trồng đất:

Những cây này không bị mất sức nhiều nên ta không cần hồi sức cho mai như mai chưng trong nhà, nhưng sau khi chưng Tết xong ta phải lặt bỏ các hoa dù đã nở hay chưa nở, vì mai quen nắng nên cây trồng chậu chưng ngoài nắng không cần phải đem vào mát.

Tỉa cành cây:Tỉa cành cây mai chậm nhất cho đến 20 âm lịch (trước ngày 15 thì tốt hơn).Tuỳ theo hình dạng của cây ta có cách tỉa thích hợp nhưng thông thường các cây mai tỉa theo dáng cây thông (trên ngắn- dưới dài để cây có hình nón), bình thường các cành được cắt tỉa đi một phần ba.

Dùng khoảng 4g urê (1 muỗng cafe nhỏ) pha với 10 lít nước tưới gốc cây và phun lên cả cây. Nếu thấy cây hồi sức lại (lá xanh hơn, tược non phát triển) thì không cần phải phun thuốc kích thích chồi lá nữa. Trường hợp cây có vấn đề thì mới sử dụng thuốc kích thích phun với liều lượng ít hơn liều lược được hướng dẫn. Nếu cành không phát triển nhiều có thể dùng một gói GA3 (1g) pha từ 30-40 lít nước phun đều lên cây và tưới gốc.

Khi cây lợi sức thì đưa từ từ ra nắng để cây quen dần, nó sẽ phát triển chồi, lá rất nhanh. Chú ý rằng đây là giai đoạn bọ trĩ và nấm hồng hoạt động (lá non, trời nắng nóng) nên pha chung hai loại thuốc có hoạt chất Hexaconazole (Anvil) Fipronil (Regent) phun lần đầu khi tỉa cành khoảng 10 ngày, phun lần thứ hai khi cây vừ nhú tước và lần sau khi lá vừa già.

Nếu năm bình thường thì việc tỉa tán cho mai nên thực hiện vào khoảng từ ngày 10 đến 20/, nếu năm nhuận thì tỉa tán muộn hơn. Các bạn chú ý việc tỉa tán rất quan trọng vì nó sẽ tạo lại sáng, tán lá cho cây. Khi cành bị cắt đi tược non sẽ phát triển thành cành mới và mang theo cả chồi trên nách lá, chồi này có thể biến thành tược hoặc nụ (phụ thuộc vào quang kỳ, phân bón, nhiệt độ và một số yếu tố khác).

Những cây không tỉa cành chắc chắn sẽ không cho hoa nhiều bằng các cây tỉa cành ngay từ đầu năm và thường dễ bị nấm bệnh. Trong việc tỉa cành các bạn chú ý rằng, cành tỉa càng gần thân thì tược phát triển càng mạnh

Vệ sinh cây: Tỉa cành cho cây xong, việc vệ sinh cho cây thực hiện rất dễ, có thể dùng vòi nước mạnh phun cho tróc bớt rong rêu nấm mốc trên cây nếu ít. Với cây có rong rêu nấm móc nhiều hơn có thể dùng Ure pha thật đặc phun vào nơi có nấm mốc (dùng nhưa cột dưới che dưới gốc không để Urê chảy xuống gốc) chừng 10 phút, sau đó dùng bàn chải chà thật mạnh cây sẽ tróc hết (nếu có máy rửa xe chỉnh áp lực thấp phun thì sạch nhất, cả những nơi như nách cành, khe kín).

Việc thay đất cho cây: Theo kinh nghiệm của các nhà vườn thì việc thay đất cho cây trong điều kiện cây bị yếu khi vừa cho hoa, kế đến trời miền nam thường nóng sau Tết nên việc thay đất không có lợi, có khi nắng nhiều mà bộ rễ bị tổn thương cây không hấp thụ đủ nước và muối khoáng nên có thể yếu đi hoặc nặng hơn có thể chết. Vì thế họ thường bón một ít phân cho cây để cây phát triển bộ rễ . Phân bón cây lúc trời nắng nóng cần thiết phải có đạm và kali, có thể dùng NPK ( 20-16-8 ) nhưng tốt nhất là phân hữu cơ như phân cá + phân bánh dầu ngâm + một ít kali (KCl). Cây bình thường, không sâu bệnh thì trong tháng 2 và đầu tháng 3 âm lịch cây sẽ có bộ lá phát triển hoàn chỉnh.

Việc thay chậu, thay đất mỗi năm không cần thiết lắm, nếu bộ rễ chưa phát triển quá nhiều trong chậu thì không cần thay đất, việc thay đất chỉ nên thực hiện sớm nhất là hai năm hoặc ba năm. Còn thay chậu chỉ nên làm khi chậu quá nhỏ so với bộ rễ của cây (thường thì đường kính chậu không quá một phần ba tán lá). Xin nhắc lại là bọ trĩ phá rất dữ trong mùa nắng nóng, nếu để cây bị phá nhiều, lá không quang hợp tốt thì năng suất hoa sẽ không cao.

Cuối tháng 3, đầu tháng 4 âm lịch miền nam bắt đầu có cơn mưa vào mùa. Nếu cần thay đất thì việc thay đất cho cây vào gần cuối tháng 3 hay đầu tháng 4 âm lịch là hợp lý. Cây trồng trong chậu nếu chất trồng chủ yếu bằng tro, xơ dừa thì mang cây ra khỏi chậu dễ dàng nếu rễ đã phát triển đầy chậu. Dùng dao thật bén gọt bỏ phần rễ già (có màu vàng sậm, khô), dùng bay nhỏ hoặc cây nhọn cạy bỏ bớt đi một phần đất trồng lâu năm.

Trước khi đưa mai vào chậu trở lại cần kiểm tra lại các lổ thoát nước, dưới đáy chậu cần lót miếng mảnh vỡ của chậu cho dễ thoát nước, trên đó là lớp cát to, một lớp phân hữu cơ, phủ lên trên một lớp chất trồng rồi mới đặt lại cây vào (không để rễ tiếp xúc trực tiếp với phân ít nhất vài tháng ), bổ sung cho đủ chất trồng, ém chặt cho cây cứng gốc. Cần chú ý là lớp phân hữu cơ và phân tro sẽ phân huỷ một thời gian cây sẽ bị lún xuống, vì thế nên đặt cây có gốc cao hơn mặt chậu để khi ổn định ta có vị trí cây như ý.

Nếu mai có chất trồng chủ yếu là đất thịt thì việc thay đất khó hơn, các bạn phải dùng bay moi quanh góc sát chậu hơn nửa vòng rồi lắc mạnh, mai sẽ tách khỏi chậu và có thể mang ra cắt bớt rễ, thay đất cho cây. Thông thường, nếu cây lớn trồng trong chậu nhỏ và nhất là trồng bằng tro trấu-xơ dừa thì mỗi năm phải thay đất 1 lần (khi thấy rễ mai bám đầy cả thành chậu). Trường hợp trồng bằng đất thịt rễ chưa bám nhiều có thể hai năm hoặc 3 năm thay cũng được (xem sự phát triển của cây).

Tuyệt đối không bón phân khi vừa thay đất vì bộ rễ không thể hấp thụ được phân, thậm chí phân có thể làm hỏng bộ rễ. Với số phân bón lót hoặc phun phân bón lá vô cơ một ít cũng đủ cho mai phát triển trong đầu mùa mưa, cộng với những cơn mưa đầu mùa, khí trời mát hẳn, sấm sét tổng hợp chất đạm tự nhiên trong không khí và đất làm cây phát triển mạnh hơn.

Công việc chăm sóc mai sau Tết coi như hoàn chỉnh. Làm các việc trên các bạn đã chuẩn bị thật tốt cho cây mai để nó tích luỹ chất dinh dưỡng trong mùa mưa, tạo nụ hoa để cho những hoa thật đẹp vào Tết tới.

Cách trồng hoa mai vàng Kỹ thuật chăm sóc hoa mai ra hoa đúng tết Hướng dẫn trồng hoa mai vàng Chăm sóc hoa mai ngày tết Chọn hoa mai ngày Tết đón may mắn Kỹ thuật trồng cây mai tứ quý Cách chăm sóc cây hoa mai hoàng yến ra hoa đẹp nhất Cách chăm sóc hoa mai ra hoa tuyệt đẹp bằng kinh nghiệm chuẩn

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