Diagnostic Protocol for Guignardia bidwellii (Black rot on

<strong>Diagnostic</strong> <strong>P<strong>rot</strong>ocol</strong> <strong>for</strong> <strong>Guignardia</strong> <strong>bidwellii</strong> 

(<strong>Black</strong> <strong>rot</strong> on grapevine) 

PEST STATUS Not present in Australia 

PROTOCOL NUMBER NDP 13 

VERSION NUMBER V1.0 

PROTOCOL STATUS Endorsed 

ISSUE DATE November 2011 

REVIEW DATE 2014 

ISSUED BY SPHDS

Prepared <strong>for</strong> the Subcommittee on Plant Health 

<strong>Diagnostic</strong> Standards (SPHDS) 

This version of the National <strong>Diagnostic</strong> <strong>P<strong>rot</strong>ocol</strong> (NDP) <strong>for</strong> <strong>Guignardia</strong> <strong>bidwellii</strong> (<strong>Black</strong> <strong>rot</strong> on 

grapevine) is current as at the date contained in the version control box on the front of this 

document. 

NDPs are updated every 3 years or be<strong>for</strong>e this time if required (i.e. when new techniques 

become available). 

The most current version of this document is available from the SPHDS website: 

http://www.padil.gov.au/Sphds

Table of Contents 

1 Introduction ................................................................................................................... 1 

1.1 Host range ............................................................................................................... 1 

1.1.1 Primary hosts .................................................................................................... 1 

1.1.2 Alternative hosts ............................................................................................... 1 

2 Taxonomic in<strong>for</strong>mation ................................................................................................. 1 

2.1 Classification ........................................................................................................... 1 

2.2 Name and Synonyms .............................................................................................. 1 

2.2.1 Synonyms ......................................................................................................... 1 

3 Detection ........................................................................................................................ 2 

3.1 Leaf, stem and fruit symptoms ................................................................................. 2 

3.2 Confusion with other diseases ................................................................................. 5 

3.3 Sampling and detection methods ............................................................................. 6 

3.3.1 Incubation of symptomatic plant material .......................................................... 6 

3.3.2 Culturing on agar media ................................................................................... 7 

4 Identification .................................................................................................................. 9 

4.1 Culture growth ......................................................................................................... 9 

4.2 Identification of conidia .......................................................................................... 11 

4.2.1 Microscopic description .................................................................................. 11 

5 Contact points <strong>for</strong> further in<strong>for</strong>mation ....................................................................... 13 

6 Acknowledgements ..................................................................................................... 13 

7 References ................................................................................................................... 14 

7.1 Other useful references ......................................................................................... 14 

8 Appendix ...................................................................................................................... 16 

Disease cycle ................................................................................................................... 16

1 Introduction 

<strong>Black</strong> <strong>rot</strong> of grape is caused by the ascomycete fungus <strong>Guignardia</strong> <strong>bidwellii</strong> and can lead to 

substantial yield losses in humid regions when it is not effectively managed. It has been 

described as one of the most economically important and destructive diseases of grapes. 

Most cultivars of Vitis vinifera as well as French/American hybrids and American bunch 

grapes are susceptible, while varieties of muscadine range in disease susceptibility from 

resistant to very susceptible. 

The fungus may be spread on grapevine cuttings and fruit. 

1.1 Host range 

1.1.1 Primary hosts 

Vitis vinifera Domestic grape 

Vitis arizonica Canyon grape 

Vitis labrusca American grape 

Vitis <strong>rot</strong>undifolia Muscadine grape 

1.1.2 Alternative hosts 

Vitis amurensis Amur grape 

Ampelopsis Wild grape 

Cissus Ornamental vine 

Parthenocissus quinquefolia Virginia creeper 

Parthenocissus tricuspidata Boston ivy 

Asplenium nidus Birds nest fern 

2 Taxonomic in<strong>for</strong>mation 

2.1 Classification 

Class: Dothideomycetes 

Order Incertae sedis 

Family: Botryosphaeriaceae 

Genus: <strong>Guignardia</strong> 

Species: <strong>bidwellii</strong> 

2.2 Name and Synonyms 

<strong>Guignardia</strong> <strong>bidwellii</strong> (Ellis) Viala & Ravaz, Bulletin de la Société Mycologique de France 8: 

63, 1892 

2.2.1 Synonyms 

Sphaeria bidwelli Ellis 1880 

Physalospora bidwelli (Ellis) Sacc., 1882 

Laestadia bidwelli (Ellis) Viala & Ravaz, 1888 

Sphaeralla <strong>bidwellii</strong> (Ellis) Ellis 1890 

Carlia <strong>bidwellii</strong> (Ellis) Magnus 1892 

Phyllachorella <strong>bidwellii</strong> (Ellis) Theiss. 1919 

Botryosphaeria <strong>bidwellii</strong> (Ellis) Petr. 1958 

Carlia <strong>bidwellii</strong> (Ellis) Prunet, 1989 

NDP Guignadia black <strong>rot</strong> 

1

3 Detection 

<strong>Guignardia</strong> <strong>bidwellii</strong> can be readily identified by macroscopic symptoms on most parts of the 

plant. The disease cycle of black <strong>rot</strong> is described in Appendix 1. 

3.1 Leaf, stem and fruit symptoms 

G. <strong>bidwellii</strong> is most likely to harbour on leaves, stems and fruit of grapevines and minor 

hosts. Sources could include imported fruit or cuttings. 

In the vineyard, symptoms are visually most evident on leaves in the spring; leaves, stems 

and fruit in the summer and on stems and fruit in autumn and winter. 

Infected leaves can be easily detected by circular lesions which vary in size but are typically 

small under field conditions (Figure 1a,d) and appear brown or tan with reddish margins and 

they characteristically contain small black pycnidia (Figure 1a-c). When inspecting vines, leaf 

lesions may be observed associated with mummified berries from the previous or current 

seasons (Figure 1d). 

a b 

c d 

Figure 1. Symptoms of G. <strong>bidwellii</strong> on leaves of Vitis vinifera (cv. Riesling); (a) on the vine, (b) on the 

laboratory bench, (c) under the dissecting microscope with pycnidia evident and (d) leaf lesions 

caused by conidia ejected from infected berries above. (Photos by M. Sosnowski, SARDI) 


2

On shoots, petioles and tendrils the lesions are initially tan, brown turning purple to black, 

sunken, elliptical to elongated and contain pycnidia or pseudothecia observed as small black 

dots (Figure 2). The bark may split and stem infections remain localised and do not usually 

extend more than several centimetres. 

a b 

d 

Figure 2. Symptoms of G. <strong>bidwellii</strong> on stems and petioles of Vitis vinifera (cv. Riesling); (a) on young 

green shoots on the vine, (b) on the laboratory bench, (c) under the dissecting microscope and (d) on 

older lignified shoots of a Vitis interspecific hybrid (cv. Vignoles) on the vine. (Photos a-c by M. 

Sosnowski, SARDI and photo d by W. Wilcox, Cornell University) 


3 

c

On the developing fruit, small, pale dots (approx. 1mm diam.) appear, rapidly becoming 

surrounded by a widening brown ring. If lesion expansion is halted due to an application of 

some fungicides or the development of age-related resistance in half grown berries, palebrown 

coloured spots with a dark ring (bird's eye effect) and a sunken centre, about 6mm 

diam, may result (Figure 3a). More typically, however, a chocolate-brown lesion expands 

through the berry (Figure 3b,c) until it becomes completely <strong>rot</strong>ted. Berries then shrivel and 

turn dark brown with numerous black pseudothecia or pycnidia developing over the surface 

(Figure 3d). Eventually, the fruit becomes dry and shrivelled, turning into hard, blue-black 

mummies that often remain firmly attached to the pedicel, although some may be shed from 

the vine (Figure 3e). 

a b 

d 

Figure 3. Symptoms of G. <strong>bidwellii</strong> on grape berries of Vitis labrusca (cv. Concord); (a-c) progression 

from small lesions to mummified berries, (d) pseudothecia on surface of mummified berry under 

dissecting microscope and (e) mummified berries. (Photos a, b, d and e by M. Sosnowski, SARDI and 

c by W. Wilcox, Cornell University) 


4 

e 

c

3.2 Confusion with other diseases 

Symptoms can be confused with those of black spot (anthracnose) disease caused by the 

fungus Elsinoe ampelina which is endemic in Australia. Leaf lesions are similar when viewed 

from a distance, but do not contain the small black pycnidia typically found in black <strong>rot</strong> 

lesions. Stems and petioles develop raised cankers with sunken centres, whereas black <strong>rot</strong> 

lesions are less “three-dimensional”. Infected fruit develop similar lesions initially, but those 

of black spot do not usually expand to include the entire berry nor evolve into mummified fruit 

(Figure 4). 

a b c 

Figure 4. Symptoms of black spot (anthracnose) disease caused by the fungus Elsinoe ampelina on 

Vitis vinifera (table grape cv. Red globe); (a) leaf lesions, (b) stem cankers and (c) berry lesions. 

(Photos by M. Sosnowski, SARDI) 

Symptoms of Phomopsis cane and leaf spot, which is caused by the fungus Phomopsis 

viticola, may also be confused with those of black <strong>rot</strong> (Figure 5). Grapevines affected by 

Phomopsis develop leaf lesions that are smaller than those due to black <strong>rot</strong>; they are often 

surrounded with a translucent halo, and do not contain pycnidia. Stem lesions are sometimes 

smaller in size, but when larger, the two diseases are very difficult to distinguish on the basis 

of lesions on the green shoots. Lesions on lignified canes often have a bleached appearance 

and develop into larger basal cankers. Fruit symptoms can be similar to black <strong>rot</strong>, with 

shrivelled mummified berries; however, this phase of the disease does not usually occur with 

Phomopsis until shortly be<strong>for</strong>e harvest, whereas it typically occurs by veraison with black <strong>rot</strong>. 

There is also a complex of pathogens including Botrytis cinerea, Aspergillus niger, Alternaria 

spp., Cladosporium spp., Rhizopus arrhizus, Penicillium spp. which may cause fruit <strong>rot</strong> of 

grapevines so should also be considered when diagnosing black <strong>rot</strong>. 


5

a b 

c d e 

Figure 5. Symptoms of Phomopsis cane and leaf spot disease caused by the fungus Phomopsis 

viticola on Vitis vinifera (cv. unknown); (a) small leaf lesions with halos, (b) green stem lesions, (c) 

bleached cane, (d) basal stem canker and (e) bunch symptoms. (Photos a-d by B. Rawnsley, SARDI 

and e by R. Emmett, DPI Victoria) 

3.3 Sampling and detection methods 

As visual symptoms can be found on leaves, stems and fruit of grapevines, they should all be 

targeted during any inspections of vineyards, post-entry quarantine or examination of 

grapevine material at the border. In the vineyard, symptoms are likely to be most obvious on 

leaves first, but mummified berries could also be present from infection in the previous 

season. Symptoms to look <strong>for</strong> are brown circular lesions (often containing small black 

pycnidia) with reddish margins on leaves; brown/purple elliptical or elongated lesions on 

stems; and brown circular lesions on fruit or shrivelled berries (details in section 3.1). In 

vineyards, symptoms can be present both during the growing season and dormancy, as 

infected, mummified berries often remain hanging on the vine since they are not detached 

from the rachis as easily as healthy berries that merely shrivel on the vine through normal 

dehydration if not harvested at maturity. 

Symptomatic leaf, stem and fruit samples should be sealed in a polyethylene bag and 

transported to a diagnostic facility, keeping samples cool during the entire process. For 

national guidelines on response to an emergency plant pest refer to PLANTPLAN (PHA 

2009). 

3.3.1 Incubation of symptomatic plant material 

Equipment: 

Dissecting microscope 

Scalpel 

Trays, polyethylene bags and paper towel 

Slides and coverslips 

Sterile distilled water (SDW) 


6

Method: 

Symptomatic leaf, stem and fruit material should be placed in a tray lined with wet paper 

towel and sealed in a plastic bag <strong>for</strong> incubation under high humidity. The samples can be left 

on the bench in normal laboratory conditions. Within 1-3 days, pycnidia on lesions may 

exude cirrhi or tendrils containing conidia which can be viewed under the dissecting 

microscope (Figure 6). 

a b 

Figure 6. Pycnidia of <strong>Guignardia</strong> <strong>bidwellii</strong> exuding cirrhi on the leaf surface of grapevine cv. Riesling 

following incubation at high humidity overnight. (Photos by M. Sosnowski, SARDI) 

Using a scalpel, scrape cirrhi from leaf surface and mount on slide in a drop of SDW. If no 

cirrhi appear then remove several pycnidia from the lesion and mount in a drop of SDW on a 

slide and crush the fruiting bodies with the side of the scalpel blade. Conidia can be viewed 

under a compound microscope and identification made as described in section 4.2. 

3.3.2 Culturing on agar media 

Equipment: 

Autoclave 

Sterile plastic petri dishes 

Laminar flow cabinet 

Incubator (25ºC, white fluorescent lights) 

Scalpel and <strong>for</strong>ceps 

Bunsen burner and alcohol 

Sodium hypochlorite solution (0.5%) 

Sterile distilled water (SDW) 

Sterile filter paper 

Parafilm 

Media preparation: 

Potato Dextrose Agar – half-strength (½ PDA) Hoffman et al. (2002) 

Potato Dextrose Agar (PDA) 9.8 g 

Bacto agar (granulated agarose) 3.5 g 

Distilled water 500 ml 

Mix ingredients and autoclave at 121ºC <strong>for</strong> 15 minutes, mixing well be<strong>for</strong>e pouring into sterile 

plastic petri dishes. 


7

Malt Extract Agar (MEA) Jailloux (1992) 

Malt Extract Agar (MEA) 16.8 g 


Mix ingredients and autoclave at 121ºC <strong>for</strong> 15 minutes, mixing well be<strong>for</strong>e pouring into sterile 

plastic petri dishes. 

Oatmeal Agar (OA) Jailloux (1992) 

Oat flakes 20 g 

Bacto agar (granulated agarose) 7.5 g 


Grind oat flakes to a powder in blender. Mix ingredients and autoclave at 121ºC <strong>for</strong> 15 

minutes, mixing well be<strong>for</strong>e pouring into sterile plastic petri dishes. 

Alternatively, use commercial <strong>for</strong>mulation of oatmeal agar. 

Method: 

In the laminar flow cabinet, remove sections of leaf, stem and fruit samples with lesions using 

a scalpel. Surface sterilise sections in 0.5% sodium hypochlorite <strong>for</strong> 60 s, rinse in sterile 

distilled water (SDW) and place on sterile filter paper to dry. Excise small pieces (approx. 3 

mm x 3 mm) from lesions and place onto surface of agar media. 

Alternatively, cirrhi exuding from pycnidia on lesions incubated as described in section 3.3.1 

can be carefully removed with a flame-sterilised scalpel and placed directly onto agar 

surface. 

Seal plates with parafilm strips and place into incubator at 25ºC under continuous fluorescent 

light. Check plates <strong>for</strong> contamination at regular intervals and subculture putative colonies of 

G. <strong>bidwellii</strong> if necessary. Pure cultures should be incubated <strong>for</strong> 15-20 days in order to induce 

sporulation. 


8

4 Identification 

<strong>Guignardia</strong> <strong>bidwellii</strong> can be readily identified by macroscopic symptoms (Section 3.1) and 

then confirmed by morphological characteristics of cultures and conidia. No molecular 

methods have been reported <strong>for</strong> the identification of G. <strong>bidwellii</strong>. 

4.1 Culture growth 

Half-strength PDA is the optimal medium <strong>for</strong> identification and conidia production (Figure 7). 

Mycelium growth rate ranges from 1-2 mm/day and appears speckled and irregular. Pycnidia 

<strong>for</strong>mation can occur within 7 days <strong>for</strong> some isolates and after 15 days all isolates produce 

fruiting bodies, from which conidia can be harvested. 

a b 

a 

c d 

e f 

Figure 7. <strong>Guignardia</strong> <strong>bidwellii</strong> cultures on ½ Potato Dextrose Agar; (a) 15 days and (b-f) 20 days of 

incubation at 25ºC under continuous fluorescent light. (e & f) Under a dissecting microscope, conidia 

are observed oozing from pycnidia. (Photos by M. Sosnowski, SARDI) 


9

Cultures grow at approx 0.5 mm/day on Malt Extract Agar (Figure 8) in a circular pattern; 

pycnidia develop within 7-15 days and are densely distributed, although they yield fewer 

conidia than on ½ PDA. Cultures grow at 1-2 mm/day on Oatmeal Agar (Figure 9) and are 

circular with a distinctive dark green, creamy appearance; few, if any, pycnidia are produced. 

a b 

Figure 8. <strong>Guignardia</strong> <strong>bidwellii</strong> cultures on Malt Extract Agar; (a) 15 days and (b) 20 days of incubation 

at 25ºC under continuous fluorescent light. (Photos by M. Sosnowski, SARDI) 

a b 

Figure 9. <strong>Guignardia</strong> <strong>bidwellii</strong> cultures on Oatmeal Agar; (a) 15 days and (b) 20 days of incubation at 

25ºC under continuous fluorescent light. (Photos by M. Sosnowski, SARDI) 


10

4.2 Identification of conidia 

Conidial ooze can be scraped from pycnidia on agar cultures or leaf lesions using a scalpel, 

and mounted on a glass slide in a drop of SDW with a cover slip placed on top. 

Viewed under a compound light microscope, conidia appear as one celled, hyaline, broadly 

ovoid, ellipsoidal or almost globose, somewhat clavate when young and slightly indented; 

spores are 5-12 × 4-7 µm in size (Figure 10). 

a b 

Figure 10. Pycnidia (a) and conidia (a&b) of <strong>Guignardia</strong> <strong>bidwellii</strong> from cultures on ½ PDA incubated 

<strong>for</strong> 15 days at 25ºC under continuous fluorescent light. (Photos by M. Sosnowski, SARDI) 

4.2.1 Microscopic description 

The following description is taken from Sivanesan and Holliday (1981). 

Pseudothecia <strong>for</strong>med as locules in a stroma, depressed globose, immersed subepidermal, 

70-180µm broad with a flat or papillate ostiolar apex (Figure 11A). The pseudothecial wall is 

made up of pseudoparenchymatic cells. Asci arise from a cushion shaped hyaline tissue at 

the base, cylindrical to clavate, 45-65 × 9-14 µm (Figure 11B). Ascospores hyaline, one 

celled, ovoid to ellipsoid, 12-17 × 6-7.5 µm, often with hyaline, mucilaginous, apical caps. 

Pycnidia mostly epiphyllous, solitary, unilocular, globose or depressed globose, with a flat or 

inconspicuous papillate ostiolar apex, 120-230 µm broad (Figure 11C). Stroma poorly 

developed on leaves but well developed on fruits. Conidiogenous cells conical to cylindrical 

(Figure 11D). Conidia one celled, hyaline, broadly ovoid, ellipsoidal or almost globose, 

somewhat clavate when young and slightly indented, 5-12 × 4-7 µm, surrounded by a 

mucilaginous sheath and with an apical hyaline appendage as long as the conidium. 

Spermatia hyaline, unicellular, rod shaped 4-7 × 0.5-2 µm. 


11

Figure 11. Microscopic description of <strong>Guignardia</strong> <strong>bidwellii</strong>; A. pseudothecium, B. ascus and 

ascospores, C. pycnidium and D. conidiogenous cells and conidia (Sivanesan and Holliday 1981). 


12

5 Contact points <strong>for</strong> further in<strong>for</strong>mation 

Dr Mark Sosnowski 

South Australian Research and Development Institute 

GPO Box 397 South Australia 5001 

Ph: 08 8303 9489 

Fax: 08 8303 9393 

Email: mark.sosnowski@sa.gov.au 

Dr Gary Kong 

Plant Science, Primary Industries and Fisheries 

Department of Employment, Economic Development and Innovation 

PO Box 102 Toowoomba Qld 4350 

Ph: 07 4688 1319 

Mob: 0428103521 

Email: Gary.Kong@deedi.qld.gov.au 

Prof Wayne Wilcox 

Cornell University 

New York State Agricultural Experiment Station 

PO Box 462 Geneva, NY 14456 USA 

Ph: +1 315 787 2335 

Email: wfw1@cornell.edu 

6 Acknowledgements 

This document is based on In<strong>for</strong>mation taken from PaDIL - Plant Biosecurity Toolbox which 

was originally authored by Gary Kong. <strong>Diagnostic</strong> methods <strong>for</strong> <strong>Black</strong> Rot of Grapes - 

<strong>Guignardia</strong> <strong>bidwellii</strong> [http://www.padil.gov.au/pbt] accessed 5 June 2009. 

The in<strong>for</strong>mation was reviewed and written into the current p<strong>rot</strong>ocol by Dr Mark Sosnowski <strong>for</strong> 

with funding from the Department of Agriculture, Fisheries and Forestry (DAFF) through the 

<strong>Diagnostic</strong> Training Scholarship programme. 

Wayne Wilcox and Judy Burr (Cornell University, NY, USA) <strong>for</strong> assistance with training and 

providing details <strong>for</strong> the diagnostic p<strong>rot</strong>ocol. 


13

7 References 

Ferrin DM and Ramsdell DC (1977) Ascospore dispersal and infection of grapes by 

<strong>Guignardia</strong> <strong>bidwellii</strong>, causal agent of grape black <strong>rot</strong> disease. Phytopathology, 67: 

1501-1505. 

Ferrin DM and Ramsdell DC (1978) Influence of conidia dispersal and environment on 

infection of grape by <strong>Guignardia</strong> <strong>bidwellii</strong>. Phytopathology, 68: 892-895. 

Hoffman LE and Wilcox WF (2002) Utilizing epidemiological investigations to optimize 

management of grape black <strong>rot</strong>. Phytopathology, 92: 676-680. 

Hoffman LE, Wilcox WF, Gadoury DA and Seem RC (2002) Influence of grape berry age on 

susceptibility to <strong>Guignardia</strong> <strong>bidwellii</strong> and its incubation period length. Phytopathology, 

92: 1068-1076. 

Jailloux F (1992) In vitro production of the teleomorph of <strong>Guignardia</strong> <strong>bidwellii</strong>, causal agent of 

black <strong>rot</strong> of grapevine. Canadian Journal of Botany, 70: 254-257. 

Pezet R and Jermini M (1989) <strong>Black</strong> <strong>rot</strong> of grapevine: symptoms, epidemiology and control. 

Revue Suisse de Viticulture, d'Arboriculture et d'Horticulture, 21: 27-34. 

PHA (2009) PLANTPLAN: Australian Emergency Plant Pest Response Plan. Version 1 May 

2009, Plant Health Australia, Canberra ACT [http://www.planthealthaustralia.com.au/] 

Ramsdell DC and Milholland RD, (1988) <strong>Black</strong> <strong>rot</strong>. In: APS, ed. Compendium of Grape 

Diseases. St Paul, USA: APS Press, 15-17. 

Sivanesan A and Holliday P (1981) CMI Descriptions of Pathogenic Fungi and Bacteria, No. 

710. Walling<strong>for</strong>d, UK: CAB International. 

Spotts RA 1977. Effect of leaf wetness duration and temperature on the infectivity of 

<strong>Guignardia</strong> <strong>bidwellii</strong> on grape leaves. Phytopathology, 67: 1378-1381. 

Wilcox W (2003) Grapes: <strong>Black</strong> <strong>rot</strong> (<strong>Guignardia</strong> <strong>bidwellii</strong> (Ellis) Viala and Ravaz.) Cornell 

Cooperative Extension Disease Identification Sheet No. 102GFSG-D4, Cornell 

University. 

7.1 Other useful references 

Alves A, Phillips AJL, Henriques I and Correia A (2007) Rapid differentiation of species of 

botryosphaeriaceae by PCR fingerprinting. Research in Microbiology, 158: 112-121. 

CAB International Distribution Maps of Plant Diseases, 1991, April (Edition 4), Map 81. 

Hoffman LE and Wilcox WF (2003) Factors influencing the efficacy of myclobutanil and 

azoxystrobin <strong>for</strong> control of grape black <strong>rot</strong>. Plant Disease, 87: 273-281. 

Hoffman LE, Wilcox WF, Gadoury DM, Seem RC and Riegel DG (2004) Integrated control of 

grape black <strong>rot</strong>: Influence of host phenology, inoculum availability, sanitation, and spray 

timing. Phytopathology, 94: 641-650. 

Janex-Favre MC, Pargueyleduc A and Jailloux F (1993) The ontogeny of pycnidia of 

<strong>Guignardia</strong> <strong>bidwellii</strong> in culture. Mycological Research, 97: 1333-1339. 

Janex-Favre MC, Parguey-Leduc A and Jailloux F (1996) The ontogeny of perithecia in 

<strong>Guignardia</strong> <strong>bidwellii</strong>. Mycological Research 100: 875-880. 

Jermini M and Gessler C (1996) Epidemiology and control of grape black <strong>rot</strong> in southern 

Switzerland. Plant Disease, 80: 322-325. 

Kummuang N, Diehl SV, Smith BJ and Graves CH (1996) Muscadine grape berry <strong>rot</strong> 

diseases in Mississippi: Disease epidemiology and crop reduction. Plant Disease, 80: 

244-247. 

Kummuang N, Smith BJ, Diehl SV and Graves CH (1996) Muscadine grape berry <strong>rot</strong> 

diseases in Mississippi: Disease identification and incidence. Plant Disease, 80: 238- 

243. 


14

Rao R and Kale SB (1965) A new species of <strong>Guignardia</strong> from India. Mycopathologia, 27: 13- 

14. 

Spotts RA (1980) Infection of grape by <strong>Guignardia</strong> <strong>bidwellii</strong> - factors affecting lesion 

development, conidial dispersal, and conidial populations on leaves. Phytopathology, 

70: 252-255. 

Wilcox WF, English-Loeb G, Dunst RM, Landers A (2006) New York and Pennsylvania Pest 

Management Guidelines <strong>for</strong> Grapes. Eds. T.H. Weigle and A.J. Muza. Cornell and 

Penn State University Cooperative Extension P 17. 


15

8 Appendix 

Disease cycle 

The black <strong>rot</strong> disease cycle is illustrated in Figure 12. The black <strong>rot</strong> fungus overwinters 

primarily in mummified fruit within the vine and on the ground, although it also can overwinter 

<strong>for</strong> at least 2 years within lesions of infected shoots that are retained as canes or spurs. 

Spring rains trigger release of ascospores (airborne sexual spores) that <strong>for</strong>m within 

mummies on the ground and in the trellis, and these can be blown <strong>for</strong> moderate distances by 

wind. Conidia (asexual spores) can also <strong>for</strong>m, both within cane lesions or on mummies that 

have remained within the trellis, and these are dispersed short distances within the vine or to 

neighbouring vines by splashing rain drops. This process can start within 1 hour of the onset 

of rain (at least 0.3 mm) and may continue <strong>for</strong> up to 8 hours after rainfall ceases (Ferrin and 

Ramsdell 1977). Infection occurs when either spore type lands on susceptible green tissue 

such as leaves, blossoms and young fruit and it remains wet <strong>for</strong> 6-24 h depending on 

temperature (Spotts 1977). 

Ascospores slowly germinate, often taking 36 to 48 hours, but eventually penetrate the 

young leaves and fruit stems (pedicels). The period of time required <strong>for</strong> symptoms to appear 

after the occurrence of infection depends on both the temperature and the age of the tissue 

at the time of infection, usually taking 8 to 25 days. The period during which these 

overwintering spores are available to cause infections depends on their source. From 

mummies on the ground, significant discharge of ascospores begins about 2 to 3 weeks after 

bud break and is virtually complete within 1 to 2 weeks after the start of flowering. In contrast, 

mummies within the trellis can continue to release both conidia and ascospores from the 

early pre-flowering period through veraison. From overwintering cane lesions, conidia can be 

dispersed from bud break through mid-summer. 

Pycnidia, produced within the lesions, continue to release conidia during wet weather 

throughout the season. The conidia will germinate and infect leaves, blossoms and young 

fruit and can cause substantial spread of the disease under warm and rainy conditions, 

particularly if berries are still susceptible to infection after conidia develop. Most berries that 

become infected near the end of their period of susceptibility do not show symptoms until at 

least 3 weeks later, and the majority do not begin to <strong>rot</strong> until 4 to 5 weeks after the infection 

event. Fruit are most susceptible to infection by the fungus from mid-flowering to about 6 

weeks after flowering, and become resistant to infection at maturity. Some older literature 

has reported that berries become resistant when they reach 5% to 8% sugar, while research 

in New York indicates that berries become resistant much earlier, 3 to 4 weeks after 

flowering. 

In summary, spore production, dispersal, infection and continued disease are favoured by 

warm, humid conditions, summer rainfall and persistent dew. . 


16

Figure 12. The disease cycle of black <strong>rot</strong>, caused by <strong>Guignardia</strong> <strong>bidwellii</strong>. (Wilcox 2003) 


17

Diagnostic Protocol for Guignardia bidwellii (Black rot on

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