Investigation of associated with Australian using cultural and molecular methods

Sung Sook Bae

A thesis submitted in fulfilment of the requirements for the degree of Doctor of Philosophy

University of New South Wales Food Science and Technology School of Chemical Engineering and Industrial Chemistry Sydney, Australia

2005 i

DECLARATION

I hereby declare that this submission is my own work and to the best of my knowledge it contains no materials previously published or written by another person, or substantial proportions of materials which have been accepted for the award of any other degree or diploma at UNSW or any other education institution, except where due acknowledgement is made in the thesis. Any contribution made to the research by others, with whom I have worked at UNSW or elsewhere, is explicitly acknowledged in the thesis.

I also declare that the intellectual content of this thesis is the product of my own work, except to the extent that assistance from others in the project’s design and conception or in style, presentation and linguistic expression is acknowledged.

Sung Sook Bae ii

ACKNOWLEDGEMENTS

I owe a tremendous debt of gratitude to numerous individuals who have contributed to the completion of this work, and I wish to thank them for their contribution.

Firstly and foremost, my sincere appreciation goes to my supervisor, Professor Graham

Fleet. He has given me his time, expertise, constant guidance and inspiration throughout my study. I also would like to thank my co-supervisor, Dr. Gillian Heard for her moral support and words of encouragement.

I am very grateful to the Australian and Wine Research Development and

Corporation (GWRDC) for providing funds for this research. Their support has truly been of great contribution to my education.

The writing of a thesis can be a lonely and isolating experience; it is obviously not possible without personal support from numerous people. Thus, I would like to give my special thanks to Ai Lin, Peter, Hugh, Mi Hwa, Lidia, Maria and Pat for making nervous and sweating moments bright and cheerful with friendship and humour. Thanks are extended to the staff of Food Science and Technology who have provided assistance and have helped to make my association with the department a rewarding experience.

I would like to express my love and gratitude to my parents, Yungsook, Sangsook, Joon

Sung, Jung Mo and Mi Hyun who have full faith in my performance and for their unconditional support. Finally, I thank my beloved husband, Minseok for his unending support and love. iii

TABLE OF CONTENTS

DECLARATION ····························································································· i ACKNOWLEDGEMENTS ·············································································ii TABLE OF CONTENTS················································································ iii ABSTRACT ··································································································vii PUBLICATIONS AND PRESENTATIONS FROM THIS THESIS ················· ix

CHAPTER 1 INTRODUCTION····································································1

CHAPTER 2 LITERATURE REVIEW 2.1 ······················································································5 2.1.1 Grapes······························································································5 2.1.2 Grape cultivation········································································5 2.2 THE PROCESS OF WINE MAKING····················································10 2.2.1 Grapes·····························································································10 2.2.2 Crushing and pre- treatments······································10 2.2.3 Alcoholic fermentation·····································································12 2.2.4 (MLF)·························································13 2.2.5 Post fermentation processes······················································ 14 2.3 ROLES OF IN ···························15 2.3.1 Fungi ·····························································································15 2.3.2 ·····························································································17 2.3.3 Bacteria ···························································································18 2.3.3.1 Lactic acid bacteria··························································18 2.3.3.2 Other bacteria····························································20 2.4 THE GRAPE SURFACE AS A MICROBIAL ECOSYSTEM ················20 2.4.1 The physical and chemical properties of grape surfaces·················22 2.4.2 Nutrients for bacterial growth on grape surfaces·····························24 2.4.3 environmental factors·······················································27 2.4.4 Agrichemical influences···································································29 2.4.5 Sources of bacterial contamination of grapes ·································31 2.4.6 Bacterial properties ···································································33 2.5 BACTERIA ASSOCIATED WITH WINE GRAPES······························35 2.5.1 Lactic acid bacteria··········································································36 iv

2.5.1.1 Isolation from ·························································36 2.5.1.2 Isolation from grapes and musts ·····································38 2.5.1.3 Isolation from environments·································40 2.5.2 bacteria ·········································································41 2.5.3 Other bacteria ··········································································43 2.6 METHODS FOR THE ANALYSIS OF BACTERIA ASSOCIATED WITH WINE GRAPES ··················································································44 2.6.1 Conventional microbiological analysis of wine grapes·····················44 2.6.2 Molecular strategies for monitoring bacterial communities··············46 2.6.3 Factors affecting the performance of molecular methods for analysis of bacteria ····································································48 2.6.3.1 DNA extraction································································48 2.6.3.2 PCR·················································································49 2.6.3.3 DGGE··············································································50 2.6.3.4 Interpretation of 16S rRNA sequence data ··················· 51 2.7 SUMMARY···························································································52

CHAPTER 3 BACTERIAL ECOLOGY OF WINE GRAPES DURING CULTIVATION AT SEVERAL IN AUSTRALIA 3.1 INTRODUCTION··················································································53 3.2 MATERIALS AND METHODS·····························································55 3.2.1 Grape samples················································································55 3.2.2 Analysis of bacteria on grapes ························································60 3.2.3 Culture on agar media ·····································································60 3.2.4 Enrichment culture···········································································62 3.2.5 DNA extraction for bacterial identification and PCR-DGGE analysis ···························································································62 3.2.6 PCR amplification of DNA for sequence identification and PCR- DGGE analysis················································································63 3.2.7 Denaturing gradient gel electrophoresis (DGGE)····························65 3.2.8 Construction of DGGE validation marker·········································66 3.2.9 DNA ·······································································67 3.2.10 Species identification········································································68 3.3 RESULTS·····························································································68 3.3.1 Climatological data for the vineyard regions in the years of 2001-2003 ·······················································································68 v

3.3.2 Population of bacteria on wine grapes during cultivation·················70 3.3.3 The diversity of bacterial species associated with wine grapes during cultivation·············································································74 3.3.3.1 Hunter Valley region, 2001-2002·····································74 3.3.3.2 Hunter Valley region, 2002-2003·····································77 3.3.3.3 Mudgee region, 2002-2003·············································87 3.3.3.4 Griffith region, 2002-2003················································92 3.3.4 Bacterial species associated with undamaged and damaged wine grapes·············································································94 3.3.5 Frequency of isolation of bacteria from wine grape samples···········99 3.4 DISCUSSION ······················································································101 3.4.1 Population and species diversity ····················································101 3.4.2 Analytical strategies for monitoring bacterial species associated with wine grapes··························································105 3.4.3 Factors affecting the bacterial ecology of wine grapes···················107

CHAPTER 4 LACTIC ACID BACTERIA ASSOCIATED WITH WINE GRAPES 4.1 INTRODUCTION·················································································112 4.2 MATERIALS AND METHODS····························································114 4.2.1 Grape samples··············································································· 114 4.2.2 Sample preparation ········································································ 114 4.2.3 Isolation of lactic acid bacteria························································ 116 4.2.4 Identification of lactic acid bacteria················································· 117 4.2.5 DNA extraction················································································ 118 4.2.6 Analysis of enrichment cultures by PCR-denaturing gradient gel electrophoresis (DGGE)································································· 119 4.2.7 Interactive growth of Oenococcus oeni, Lactobacillus kunkeei and Lactobacillus lindneri·······························································121 4.3 Results ·····························································································122 4.3.1 Isolation of lactic acid bacteria from enrichment cultures ···············122 4.3.2 Identification of isolates··································································129 4.3.3 Analysis of lactic acid bacteria in enrichment cultures by PCR-DGGE···············································································132 4.3.4 Interactive growth of O. oeni and lactic acid bacteria ·····················139 4.4 DISCUSSION ······················································································141 vi

CHAPTER 5 OCCURRENCE AND SIGNIFICANCE OF thuringiensis ON WINE GRAPES 5.1 INTRODUCTION·················································································146 5.2 MATERIALS AND METHODS ····························································148 5.2.1 Microbial strains··············································································148 5.2.2 Grape samples···············································································149 5.2.3 Isolation and identification of B. thuringiensis from grapes·············149 5.2.4 Isolation of B. thuringiensis from wines and grape ·················151 5.2.5 Interaction between B. thuringiensis and other microorganisms ····151 5.2.6 Growth and survival of B. thuringiensis in grape juice and wine ····152 5.2.7 Growth of microorganisms in mixed culture with B. thuringiensis···153 5.3 RESULTS····························································································153 5.3.1 Isolation of B. thuringiensis from commercial insecticides··············153 5.3.2 Occurrence of B. thuringiensis on grapes, juice and wine samples··························································································154 5.3.3 Interaction between B. thuringiensis and microorganisms significant in wine production 5.3.3.1 Yeasts·············································································156 5.3.3.2 Bacteria ··········································································156 5.3.3.3 Filamentous fungi···························································157 5.3.4 Survival and growth of B. thuringiensis in grape juice and wine·····159 5.3.5 Interactive growth of B. thuringiensis and S. cerevisiae ·················160 5.3.6 Interactive growth of B. thuringiensis and O. oeni ··························161 5.4 DISCUSSION ······················································································164

CHAPTER 6 CONCLUSIONS···································································169

CHAPTER 7 BIBLIOGRAPHY··································································174

APPENDIX 1 ·····························································································199 APPENDIX 2 ·····························································································202 APPENDIX 3 ·····························································································204 APPENDIX 4 ·····························································································206 APPENDIX 5 ·····························································································207 vii

ABSTRACT

This thesis presents a systematic investigation of bacterial species associated with wine grapes cultivated in Australian vineyards during 2001-2004. Grapes, sampled throughout cultivation, were analysed for bacterial species using a combination of cultural and molecular methods. Red (Shiraz, , ) and white

(, Semillon, ) grape varieties were examined. Factors affecting the bacterial ecology of grapes were considered.

The bacterial populations of mature undamaged grapes at were consistently low at 102-103 CFU/g. Higher populations (103-106 CFU/g) were found on grapes at earlier stages of maturity and correlated with application of Bacillus thuringiensis as a biological pesticide. B. thuringiensis was the most prevalent bacterial species on wine grapes throughout cultivation, as determined by plate culture, enrichment culture and

PCR-DGGE. B. thuringiensis carried over into wine processing but did not grow in juice or wine and did not adversely affect the growth of cerevisiae or

Oenococcus oeni in liquid culture. B. thuringiensis inhibited the growth of several spoilage and mycotoxigenic fungi found on grapes.

Curtobacterium flaccumfaciens was the second most prevalent species detected on wine grapes. Its populations rarely exceeded 103-104 CFU/g. Other bacteria (Arthrobacter,

Bacillus, Microbacterium, Pantoea, Pseudomonas, Sphingomonas) were sporadically found on grapes.

Lactic acid bacteria and acetic acid bacteria were rarely detected on undamaged grapes viii by culture and PCR-DGGE methods. A greater incidence of lactic acid bacteria was detected by specific enrichment procedures, especially on damaged grape berries.

Species found were Lactobacillus plantarum, Lactobacillus mali, Lactobacillus lindneri and Lactobacillus kunkeei. The malolactic organism, O. oeni, was never isolated from any grape sample, raising questions about its enological origin. Enrichment cultures also revealed the presence of other bacteria (e.g. Sporolactobacillus inulinus, siamensis) not previously found on wine grapes.

Atypical, hot and dry conditions during cultivation may account for the low populations of bacteria found on wine grapes. This factor combined with the overwhelming presence of B. thuringiensis prevented meaningful comparisons of data to determine influences of vineyard location, grape variety, grape maturity, climate and viticultural practices on the bacterial ecology of grapes. More systematic and controlled studies of these variables are required. ix PUBLICATIONS AND PRESENTATIONS FROM THIS THESIS

Bae, S., Fleet, G.H. and Heard, G.M. (2005) Lactic acid bacteria associated with wine grapes. J. Appl. Microbiol. (accepted and under revision)

Bae, S., Fleet, G.H. and Heard, G.M. (2004) Significance and occurrence of Bacillus thuringiensis on wine grapes. Intl. J. Food Microbiol. 94, 301-312.

Fleet, G.H., Beh, A.L., Prakitchaiwattana, C.J., Bae, S. and Heard, G.M. (2002) The and bacterial ecology of wine grapes. International Union of Microbiological Societies (IUMS) for the Xth International Congress of Bacteriology and Applied , The World of Microbes, Paris 27 July-1 August.

Bae, S., Fleet, G.H. and Heard, G.M. (2003) Significance and occurrence of Bacillus thuringiensis on wine grapes. Poster paper presented at the 11 Australian Conference of AIFST. Noosaville, Queensland, 26-28 March. G 1 CHAPTER ONE INTRODUCTION

Microorganisms, representing various yeasts, bacteria and filamentous fungi, are intimately associated with the production of wines, and are major determinants of product quality and process efficiency (Fugelsang 1997; Fleet 1998). In a positive context, yeasts conduct the alcoholic fermentation of grape juice into wine, and lactic acid bacteria enhance wine flavour and stability through the malolactic fermentation.

However, microorganisms also have various negative impacts throughout the production chain. They can depreciate the quality of the grapes, contribute to sluggish, stuck or failed , and spoil the product at various stages of the process

(Fleet 1998; 2001). This thesis is concerned with the bacteria associated with wine production, in particular, the bacterial ecology of wine grapes.

The main bacteria of winemaking are species of lactic acid bacteria (Oenococcus oeni,

Lactobacillus spp., Pediococcus spp., Leuconostoc spp.), acetic acid bacteria

(Acetobacter spp., Gluconobacter spp.) and, infrequently, species of Bacillus and

Clostridium. With the exception of O. oeni and possibly Lactobacillus plantarum that are responsible for the malolactic fermentation (Henick-Kling 1993; Bartowsky 2005), these bacteria have undesirable consequences and contribute to problems of spoilage, taints and stuck/sluggish fermentations (Wibowo et al. 1985; Drysdale and Fleet 1988;

Sponholz 1993; Bisson 1999; Lonvaud-Funel 1999; du Toit and Pretorius 2000). Much research has been conducted on the occurrence, growth and significance of these bacteria within the wine and winery environments, but there is very little information about their association with wine grapes.

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Grapes are considered to be a primary source of microorganisms in the wine and winery ecosystems. In recent years, there has been significant study and debate on the origin of wine yeasts, and the importance of grapes as a source of these organisms (Martini et al.

1996; Mortimer and Polsinelli 1999; Fleet et al. 2002). Such scrutiny has not been extended to the bacteria of winemaking where, in contrast, there has been very little study in this context.

The early literature certainly shows that various species of lactic acid bacteria and acetic acid bacteria can be isolated from wine grapes (Kunkee 1967; Amerine and Kunkee

1968; Wibowo et al. 1985; Drysdale and Fleet 1988). Bacteria with malolactic activity have been demonstrated in this flora, although there are very few conclusive reports on the isolation of O. oeni from grapes. Most studies represent simple, qualitative descriptions and identification of the bacteria that were isolated. At best, it can be concluded that mature, sound grape berries may harbour low populations (101-103

CFU/g) of lactic acid bacteria and acetic acid bacteria. Damaged or spoiled grapes may have much higher populations (Fleet 1993).

The surface of grapes represents a phyllospheric habitat and, by analogy with other fruit and plant products, many factors are known to affect the survival and growth of bacteria at this location (Fleet 2003b). Such factors include climatic influences, agricultural

(viticultural) practices, and damage to the fruit. Moreover, the diversity and populations of bacterial species associated with grapes could change and evolve as the grape berry develops and matures on the vine, and there may be influences of grape cultivar. The bacterial species associated with the grape surface are not only significant as a source of winery bacteria but, through natural microbial interactions, they could also influence the

G G 3 diversity and population of yeasts and filamentous fungi that develop on grapes

(Belanger and Avis 2002; Fleet 2003a). In this context, they could impact on grape quality and the yeast flora that enter the winery environment.

In summary, while much research has been devoted to describing the occurrence and growth of bacteria in wines, there is little information about the diversity of bacterial species present on grapes and the factors that affect this bacterial community. Research is needed to determine what bacterial populations occur on wine grapes and what factors affect their occurrence and survival. Such information and understanding should enable practices that enhance grape quality, allow better management of the malolactic fermentation and minimize the risks of problem fermentations and wine spoilage.

PROJECT OBJECTIVES

The overall goal of this research project is to undertake a systematic investigation of bacterial species associated with wine grapes cultivated at several Australian vineyards.

A combination of standard cultural methods and newer molecular methods will be used to analyse these bacteria.

Specific aims are to:

z Investigate and identify the bacterial community associated with the surfaces of

wine grapes.

z Correlate, where possible, the occurrence and populations of bacterial species

on grapes with viticulture practices.

Factors to be considered include vineyard location, temperature and rainfall,

grape cultivar, grape berry maturity, physical damage to the grape berry, and

G G 4

application of agrichemicals.

z Investigate any effects (antagonistic, stimulatory) of dominant bacterial isolates

on yeasts, filamentous fungi and other bacteria associated with grapes and wine

production.

G G 5 CHAPTER TWO LITERATURE REVIEW

This thesis is concerned with bacteria associated with wine grapes and the application of molecular methods to their analysis. Key studies associated with these topics will be presented in the Introduction and Discussion sections of subsequent experimental

Chapters. This Chapter will provide a brief background to the role of bacteria in winemaking followed by a more detailed review of the literature describing the bacteria on wine grapes. A final section of this Chapter will provide some background information on molecular methods used to identify and analyse bacteria in food ecosystems.

2.1 VITICULTURE

2.1.1 Grapes

Viticulture is the study of grapes and grape cultivation for wine production. Numerous varieties of the grape, vinifera, are used in winemaking to make both white and red wines, with the particular variety determining the fruity or floral characteristics of the wine product. Some main grape varieties of are Chardonnay, ,

Sauvignon Blanc, Semillon, Traminer, and Müller-Thurgau, while those used to make red wines include Cabernet Sauvignon, , , Shiraz, ,

Grenache, and (Dry and Gregory 1992; Boulton et al. 1995; Fleet 2001).

2.1.2 Grape cultivation

Most grape varieties are usually grown in a nursery for one year to produce rooting, and

G G 6 then they are planted in the vineyard. Grapevines require three to six years after planting to reach full economic production. A considerable amount of viticultural research has identified strategies for optimizing the cultivation of grapes for winemaking. These strategies include pruning, and canopy management of the vine, water supply and irrigation, dispersal of drainage water, soil management for optimal growth of grape roots, and pest and disease control. There is extensive literature in the field of viticulture and the reader is referred to some key texts for further information (Jacquelin and

Polulain 1962; Winkler 1973; Coombe and Dry 1992; Jackson and Lombard 1993;

Ribéreau-Gayon et al. 2000).

The production of wine grapes follows an annual cycle - budburst, flowering, berry formation, veraison and harvest. The formation of leaf buds on the grapevine begins in spring. During the next 3-4 months, the vine develops, producing mature grape berries through the stages of inflorescence, flowering, pollination and fertilisation of the ovary, fruit and berry setting, growth of the green berry and ripening of the berry. More details of these developmental stages are given in Table 2.1 and pictorial illustration is given in

Fig. 2.1 (Pratt 1971; Coombe 1992, 1995; Coombe and McCarthy 1997; Kennedy 2002).

It is not known how the bacterial ecology of the grape varies throughout this development and will be examined in this thesis.

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Table 2.1 Key stages in the development of wine grapes in the vineyard

Budburst Dormancy of vine bud is broken; generally recognized as formation of a green tip or shoot, and visible formation of leaf tissue. Usually occurs in March-May in northern hemisphere and September-November in southern hemisphere Shoot and leaf development Progressive unfolding of leaves; beginning of inflorescence and flower formation Flowering Loosening and detachment of flower caps, beginning of inflorescence and flower formation Setting and development of grape berries (fruiting) Berries usually hard and green, progressively increase in size from about 2mm in diameter (peppercorn size) to about 7mm in diameter (pea size); accompanied by closure of bunches of berries. This stage lasts about 60-70 days Veraison Berries begin to soften, change colour and increase in size; sugar concentration increases, acidity decreases. Generally starts about August-September in northern hemisphere and December-January in southern hemisphere and lasts about 40-50 days Engustment Prior to harvest often late in the ripening process, sudden development/accumulation of grape flavour Harvesting Berries are harvested when fully ripe and proceed to senescence after this stage Pratt 1971; Coombe 1992, 1995; Coombe and McCarthy 1997; Fleet et al. 2002

G Shoot and inflorescence development Flowering Berry Development Ripening E-L number 1 2 3 4 5 7 9 11 12 13 14 15 16 17 18 19 20 21 23 25 26 27 29 31 32 33 3435 36 37 38

38. Harvest

27. Setting 35. Veraison 12. Shoots 10cm 31. Berries pea size 23. Full bloom

19. Flowering begins Engustment

4. Budburst

Veraison

setting

Days after flowering 0 20 40 60 80 100 120

Fig. 2.1 Adoption of a system for identifying grapevine growth stages as produced by Coombe (1995) and Kennedy (2002) 8 E-L E-L number ALL STAGES number ALL STAGES 1. Winter bud Development Berry 27. Setting young berries enlarging (>2mm diam.), 2. Budswell bunch at right angles to setem ho n nlrsec development inflorescence and Shoot 3. Woolly bud-brown wool visible 29. Berries pepper-corn size (4 mm diam.); 4. Green tips first leaf tissue visible bunches tending downwards 5. Rosette of leaf tips visible 31. Berries pea size (7 mm diam.) 7. First leaf separated from shoot tip 32. Beginning of bunch closure, berries touching 9. 2-3 leaves separated; shoots 2-4 cm long (if bunches are tight) 11. 4 leaves separated 33. Berries still hard and green 12. 5 leaves separated; shoots about 10 cm long; 34. Berries begin to soften; start increasing

inflorescence clear Ripening 35. Berries begin to colour and enlarge 13. 6 leaves separated 36. Berries with intermediate Brix value 14. 7 leaves separated 37. Berries not quite ripe 15. 8 leaves separated, shoot elongateing rapidly; Engustment single flowers in compact groups 38. Berries harvest-ripe 16. 10 leaves separated 17. 12 leaves separated; inflorescence well developed, single flowers separated 18. 14 leaves separated; flower caps still in place, but cap colour fading from green 19. About 16 leaves separated; beginning of flowering (first flower caps loosening) Flowering 20. 10% caps off 21. 30% caps off 23. 17-20 leaves separated; 50% caps off (full bloom) 25. 80% caps off 26. Cap-fall complete

Fig. 2.1 (continued) Adoption of system for identifying grapevine growth stages as produced by Coombe (1995) and Kennedy (2002) 9 G 10

2.2 THE PROCESS OF WINEMAKING

The process of winemaking varies with the type of wine being produced, the technology innovations that are applied and with the region or country. Specific details can be obtained from key texts (Boulton et al. 1995; Ribéreau-Gayon et al. 2000). However, there are some common principles and operations which are outlined in the following section as background to understanding the role of microorganisms in wine production.

Fig. 2.2 gives the main steps in the production of white and red wines.

2.2.1 Grapes

The varieties of grapes used in winemaking have been mentioned already in Section 2.1.

The grapes for wine production are harvested at an appropriate stage of maturity.

Particularly important are the concentrations of sugars and acids, which are major constituents of the juice and have an important impact on its fermentation properties

(Fleet 1998, 2001).

2.2.2 Crushing and pre-fermentation treatments

Red grapes are destemmed and crushed to yield the pulp and juice (must), which is transferred to tanks for fermentation. Fermentation can occur naturally or by inoculation with selected yeasts. For red winemaking, crushing facilitates of the grapes into a fermentable substrate, distribution of microflora of the grape surface throughout the must, and extraction of and tannin components of skin into must

(Boulton 1995). This extraction is also assisted by produced during the early stage of alcoholic fermentation. White wines are produced by the fermentation of grape juice without the solid part of the berry (skins and seeds) to prevent detrimental effects

G 11

RED WINES WHITE WINES Grapes Grapes

Crushing; addition Crushing; addition of sulphur dioxide of sulphur dioxide

Juice and Skins Pre-fermentation treatments; settling Inoculation with yeast (optional) (removal of skins) Maceration and partial fermentation (colour and tannin extraction) Juice

Pressing Inoculation with yeast (removal of skins) (optional)

Completion of Alcoholic fermentation alcoholic fermentation followed by off yeast less followed by racking off yeast

Wine Wine

Malolactic fermentation Malolactic fermentation (optional) (optional)

Aging in barrels or Aging other vessels (optional)

Fining, clarification, packaging Fining, clarification, packaging

FINAL PRODUCT FINAL PRODUCT

Fig. 2.2 Outline of process for and white wine production (Fleet 2001) G 12 on wine quality, and grape skin from colouring the must. Since aromas and aroma precursors are located in the grape skin or in the underlying cell layers in most quality cultivars (Ribéreau-Gayon et al. 2000), contact of grape juice with skin or seeds prior to fermentation is necessary and critical. Pre-fermentation operations such as varying the degrees of crushing, cold settling, pressing for juice extraction and clarification are, therefore, carefully managed to determine the final tastes of white wine

(Ewart 1995).

2.2.3 Alcoholic fermentation

The alcoholic fermentation is the primary fermentation, where yeasts transform sugars in grape juice (mainly and fructose) into ethanol (the main metabolite of wine) and . This process can be conducted either as an indigenous/wild fermentation, or as an induced /seeded fermentation. With indigenous fermentation, yeasts resident in the grape juice initiate and complete the fermentation. With seeded fermentation, selected yeast strains, generally those of or

Saccharomyces bayanus, are inoculated into the juice at initial populations of 106-107 cells/mL. The advantages and disadvantages of indigenous and seeded fermentations have been well discussed (Kunkee and Amerine 1970; Kunkee and Goswell 1977;

Kunkee 1984; Reed and Nagodawithana 1988; Kunkee and Bisson 1993; Kunkee and

Vilas 1994; Heard 1999; Pretorius 2000).

Fermentation of the juice was traditionally conducted in large wooden barrels or concrete tanks, but most modern now use sophisticated stainless steel tanks with facilities for process management. However, some premium-quality white wines

(e.g. Chardonnay) may be fermented in wooden (oak) barrels. White wines are

G G 13 generally fermented at 10-18qC for 7-14 days or more, where the lower temperature and slower fermentation rate favour the retention of desirable volatile flavour compounds.

Red wines are fermented for about 7 days at 20-30qC, where the higher temperature is necessary to extract colour from the grape skins.

Alcoholic fermentation is considered complete when the fermentable sugars of juice are completely utilized. The wine is then drained or pumped (racked) from the sediment of yeast and grape material (lees) and transferred to stainless steel tanks or wooden barrels for malolactic fermentation, if desired, and aging. Leaving the wine in contact with the lees for long periods is not encouraged because the yeast cells autolyze, with the potential of adversely affecting wine flavour and providing nutrients for the subsequent growth of spoilage bacteria (Kunkee and Amerine 1970; Fleet 1998, 2001).

2.2.4 Malolactic fermentation (MLF)

Malolactic fermentation is a secondary fermentation caused by growth of certain lactic acid bacteria. MLF may be due to malolactic bacteria naturally present in wine, but nowadays malolactic fermentation is often induced with commercial starter cultures

(van Vuuren and Dicks 1993; Costello et al. 2003). Oenococcus oeni (formerly

Leuconostoc oenos) (Dicks et al. 1995) and possibly Lactobacillus plantarum (Renault et al. 1988) are the preferred species for carrying out malolactic fermentation. One of the main effects of the MLF is a decrease in total acidity, resulting from the decarboxylation of L-malic acid to L-lactic acid. The decrease in acidity is particularly desirable for high-acid wines in cool climate regions, and gives the wine a softer and mellower taste. Moreover, MLF contributes additional metabolites, such as , , , volatile acids, and 2, 3-butanediol that may enhance the

G G 14 flavour complexity of wine (Davis et al. 1985; Bartowsky and Henschke 1995;

Lonvaud-Funel 1999; Bartowsky et al. 2002; Liu 2002). There is a perception among winemakers that wines which have not undergone MLF are microbiologically less stable, and have an increased risk of spontaneously undergoing this reaction after bottling. If this occurs, the bottled wines become turbid, gassy, show sediment, and are considered to be spoiled (Rankine and Bridson 1971). Wines with completed MLF appear to have greater microbiological stability and are less prone to spoilage by other species of bacteria. It is thought that growth of bacteria during MLF deprives substrates of wine for subsequent bacterial growth and spoilage, and produces bacteriocin as a further inhibitory factor (van Reenen et al. 1998; Bauer et al. 2005).

MLF is not necessarily beneficial to all wines (Kunkee 1991). Wines produced from grapes grown in warmer climates tend to be less acid. Further reduction in acidity by

MLF is deleterious to overall sensory balance, and also it increases their pH to values at which spoilage bacteria are more likely to grow. However, preventing the natural occurrence of malolactic fermentation in these wines is an extra technical burden.

Consequently, many winemakers prefer to encourage the malolactic fermentation and later adjust wine acidity, if necessary (Fleet 2001).

2.2.5 Post fermentation processes

Most wines are not stored for lengthy periods after completion of the fermentation. If storage is necessary, it is generally done in stainless steel tanks. Some white wines (e.g.

Chardonnay) may be aged in wooden barrels. Premium quality red wines are aged for periods of 1-2 years by storage in wooden (oak) barrels. During this time, chemical reactions occur between wine constituents and components extracted from the wood of

G G 15 the barrels and contribute to flavour development (Rankine 1989; Boulton et al. 1995).

Critical points for control during storage and aging, are exclusion of and addition of dioxide to free levels of 20-25 Pg/mL. These controls are necessary to prevent the growth of spoilage bacteria and yeasts, and unwanted oxidation reactions.

Before bottling, wines may be cold stored at 5-10qC to precipitate excess tartrate and then clarified by application of one or more processes, including the addition of fining agents (bentonite, albumen, isinglass, gelatin), centrifugation, pad filtration, and membrane filtration. For some white wines with residual sugar, sorbate up to

100-200 Pg/mL may be added to control yeast growth (Rankine 1989; Ewart 1995).

2.3 ROLES OF MICROORGANISMS IN WINEMAKING

The surface of grape berries represents a phyllospheric habitat for a diversity of fungal, yeast and bacterial species that have various impacts on the efficiency and quality of wine production.

2.3.1 Fungi

Filamentous fungi are most significant organisms that contaminate and colonise the surface of grapes during cultivation. They play an important role in the physical and chemical stability of grapes, and they may impact on the sensory properties of the future wine. The filamentous fungi of importance in wine production include species of

Alternaria, Aspergillus, Botrytis, Cladosporium, and Penicillium. These species are well known for their ability to spoil grapes prior to harvest (Pitt and Hocking 1997; Zahavi et al. 2000). They include (sour bunch rot), Penicillium spp. (blue rot),

G G 16

Aspergillus niger (black mold rot), Rhizopus nigraicans (rhizopus rot), and

Cladosporium herbarum and Alternaria tenuis (sour rot). Often, mould contamination and spoilage of grapes is accompanied by secondary growth of native yeasts and bacteria, that can add further spoilage (Fugelsang 1997). In addition, concerns regarding the presence of mycotoxin on mould-damaged grapes are continuing to receive attention, since the mycotoxin, ochratoxin A, has been found in some wines. Its occurrence has been correlated with the growth of Aspergillus carbonarius and other species on grapes during cultivation (Cabañes et al. 2002; Sage et al. 2002; Battilani et al. 2004). The presence of mycotoxins in mould-damaged grapes could be a government regulatory issue and affect the decision of whether a winery will accept or reject the supply of grapes from growers (Fugelsang 1997). In addition to spoilage of grapes, fungi are able to grow on the surface of stored, grape juice concentrate, on the outside and inside surfaces of wooden storage containers and on corks, even though they cannot grow in wine due to sensitivity to ethanol. The formation of sensorially powerful metabolites, that give corky, earthy taints may enter the wines to cause its spoilage (Lee and

Simpson 1993; Fleet 2003a).

Although fungal development, alone or associated with other microorganisms, lowers potential grape quality, B. cinerea can be unique among problematic fungi depending on the condition of infection. In certain circumstances, the development of B. cinerea on grapes results in an overripening process (referred to as ) by facilitating dehydration of free water and concentration of grape sugars and permits the production of the world’s most sought-after sweet wines (Donèche 1991, 1993; Fugelsang 1997;

Ribéreau-Gayon et al. 2000).

G G 17

2.3.2 Yeasts

It has been recognized since the studies of Louis Pasteur about 150 years ago that yeasts play a key role in wine production. Primarily, they conduct the alcoholic fermentation, but at later stages of production, they can cause spoilage (Fleet 2001).

Yeasts occur as natural flora on the surface of grapes. Consequently, grapes are a primary source of yeasts associated with wine fermentation. During alcoholic fermentation, yeasts species and strains within the genera Hanseniaspora, ,

Metschnikowia, and sometimes Kluyveromyces grow during the early stages, but eventually die off due to toxicity of the increasing concentration of ethanol, leaving

Saccharomyces cerevisiae or as the dominant species to complete the fermentation (Fleet and Heard 1993; Fleet 1998, 2001; Pretorius 2000;

Bisson and Block 2002). Saccharomyces cerevisiae and S. bayanus have become universally accepted as the principal wine yeasts. Particular strains of these yeasts have been commercialised and are now sold to winemakers as active dry cultures for inoculation into freshly crushed juice. It is assumed that the inoculated S. cerevisiae and

S. bayanus will overwhelm and diminish the growth of the indigenous flora, and dominate the fermentation. However, even in these inoculated ferments, indigenous flora will always be present and will make variable contributions to the process, depending on the competitive success of the inoculated S. cerevisiae or S. bayanus.

Even though indigenous non-Saccharomyces yeasts die off the in the early stages of fermentation, they will have previously grown to significant populations and put their imprint on wine character (Heard and Fleet 1985). Also, there will be contributions from indigenous strains of S. cerevisiae and S. bayanus (Fleet 2001).

G G 18

Yeast can spoil wines at several stages during wine production. The growth of an undesirable yeast during alcoholic fermentation, storage or bottling may cause unacceptable sensory properties of appearance, aroma and flavour. These defects include taints (Hanseniaspora uvarum during initial stages of alcoholic fermentation), film formation (species of Candida, Metschnikowia, Pichia and

Hansenula during storage), re-fermentation ( bailii,

Schizosaccharomyces pombe, S. cerevisiae during storage and after packaging), mousy off flavour (Dekkera and ) and production of haze, turbidity and volatile acidity (species of Saccharomyces and Brettanomyces in bottled wine) (Sponholz 1993; du Toit and Pretorius 2000; Grbin and Henschke 2000).

2.3.3 Bacteria

Bacteria are prominent microorganisms in wine production and contribute to wine flavour and other properties by a range of mechanisms and activities (Fleet 2003a).

While lactic acid bacteria are the main bacterial species of wine production, other bacteria also have significant roles.

2.3.3.1 Lactic acid bacteria

Lactic acid bacteria are one of the most important groups of bacteria in wine production.

Depending on the stage of the winemaking process and the species that grow, they can have positive or negative impacts on the quality of wine. They are primarily responsible for the MLF, but can also cause spoilage. The occurrence and significance of lactic acid bacteria in wines have been the subject of extensive studies that are well reviewed in the literature (Kunkee 1967, 1974; Davis et al. 1985; Wibowo et al. 1985; Henick-Kling

1993; Lonvaud-Funel 1999; Liu 2002).

G G 19

Almost 100 years ago, it was observed that many wines underwent a natural secondary fermentation about 2-4 weeks after completion of the alcoholic fermentation. This fermentation has been called the MLF and its ecology and biochemistry have been extensively studied (Kunkee 1967, 1974, 1991; Lafon-Lafourcade and Ribéreau-Gayon

1984; Henick-Kling 1993; Bartowsky and Henschke 1995; Boulton et al. 1995;

Lonvaud-Funel 1995, 1999; Bartowsky 2005). It is conducted by lactic acid bacteria that originate from the grapes and survive the alcoholic fermentation, or come from winery equipment. They generally exist at low or non-detectable levels (<10-100

CFU/mL) in the wine at the completion of alcoholic fermentation but, over 2-4 weeks, grow to populations of 107-108 CFU/mL. The most notable feature of this growth is the stoichiometric decarboxylation of L-malic acid to L-lactic acid and carbon dioxide, causing deacidification of the wine and an increase in pH by about 0.3-0.5 unit. This deacidification gives a most desired improvement in wine sensory quality, especially for high acid wines with initial pH values of 3.0-3.5. However, for less acid wines (e.g., pH

3.5-4.0), further deacidification may depreciate sensory quality and increase the probability of wine spoilage. Besides deacidification, MLF may also enhance sensory quality and microbiological stability (Bartowsky et al. 2002; Liu 2002).

Oenococcus oeni (Dicks et al. 1995) is the main species that conducts MLF, and it is uniquely found in the winery environment. Other species of lactic acid bacteria, namely

Pediococcus parvulus, Pediococcus pentosaceus, Pediococcus damnosus, and various species of Lactobacillus can also conduct the MLF but, they often give unpleasant off- flavours (Table 2.2). In addition, lactic acid bacteria which have the ability to grow up to 109 CFU/mL in the early stages of alcoholic fermentation, may cause stuck or sluggish fermentation by inhibiting the growth of wine yeast (Huang et al. 1996; Bisson

G G 20

1999; Edwards et al. 1999; du Toit and Pretorius 2000).

2.3.3.2 Other bacteria

Acetic acid bacteria (Acetobacter and Gluconobacter spp.) are also well known for spoilage of injured grapes and their ability to oxidize ethanol at the air interface or the middle of barreled wines, to give vinegary spoilage. Their early growth in grape juice could also lead to stuck or sluggish alcoholic fermentation (Drysdale and Fleet 1988,

1989a). The potential for acid-tolerant, ethanol-tolerant species of Bacillus and

Clostridium to grow in wines should not be underestimated (Gini and Vaughn 1962;

Murrell and Rankine 1979; du Toit and Pretorius 2000). Various species of Actinomyces,

Streptomyces can grow within the cracks and pores of wooden barrels and lenticels of corks, producing metabolites that give overpowering, deleterious taints when leached into the wine (Fleet 1998, 2001). The main problems caused by growth of bacteria in wine production are summarized in Table 2.2.

2.4 THE GRAPE SURFACE AS A MICROBIAL ECOSYSTEM

The surface of wine grapes can be considered as a unique phyllospheric habitat which represents the primary source of microorganisms associated with wine production. The grape surface provides an environment for microorganisms to attach and, nutrients translocated from inside the berries, allow colonization by a diversity of filamentous fungi, yeasts and bacteria. Several stages occur in the microbial colonization of grape surfaces. These are contamination, attachment and immobilization of cells to the surface, and accessing of nutrients for growth. The bacterial flora of grapes needs to be considered and studied in the context of the grape surface being a unique ecosystem.

G G 21

Table 2.2 Spoilage reactions caused by the growth of bacteria during wine production

Spoilage reaction Responsible Reference microorganisms

Acidification and stuck, sluggish Lb. kunkeei Ingledew and Kunkee 1985; fermentation Lb. brevis Sponholz 1993; Edwards et al. Production of unacceptable acetic acid and Lb. nagelii 1998a, 1998b, 2000; Bisson 1999 D-lactic acid from metabolism of sugars. O. oeni

Acrolein formation Lb. cellobiosus Davis et al, 1988; Sponholz 1993; Bitterness arising from metabolism of Lb. hilgardii Bartowsky and Henschke 1995; to acrolein Leuc. mesenteroides Fugelsang 1997 P. parvulus

Buttery flavour Lb. plantarum Martineau and Henick-Kling 1995 Due to excessive diacetyl production O. oeni Pediococcus spp.

Formation of biogenic amines Lb. hilgardii Delfini 1989; Coton et al. 1998; (e.g. histamine, tyramine, putrescine, P. damnosus, Arena and Manca de Nadra 2001; cadaverine, phenylethylamine) O. oeni Liu 2002

Geranium off-odour O. oeni Edinger and Splittstoesser 1986 Hydrogenation of sorbic acid to sorbic causing unpleasant geranium plant-like odour

Mannitol taint Lb. brevis Sponholz 1993 Enzymatic reduction of fructose to mannitol, cause estery off flavours.

Mousey taint Lb. brevis, Fugelsang 1997; Costello et al. 2001 Microbial production of acetyl- Lb. cellobiosus, tetrahydropyridines Lb. hilgardii

Production of ethyl carbamate Lb. brevis Liu and Pilone 1998; Azevedo et al. precursors Lb. buchneri 2002; Liu 2002 Lb. hilgardii O. oeni

Ropiness A. xylinum Sponholz 1993; Gindreau et al. Production of extracellular , P. damnosus 2001 cause slimy and viscous mouth feel Leuc. mesenteroides

Tartaric degradation Lb. brevis Sponholz 1993 Depreciates sensory quality Lb. plantarum Martineau and Henick-Kling 1995

Oxidation of ethanol to acetaldehyde A. aceti Drysdale and Fleet 1989b; Sponholz and acetic acid A. pasterianus, 1993 G. oxydans

Production of acidic (butyric) off- Bacillus sp. Gini and Vaughn 1962; Murrell and flavours Clostridium sp. Rankine 1979; Sponholz 1993

Earthy, corky taints Actinomyces Fleet 1998, 2001 Streptomyces A, Acetobacter; G, Gluconobacter; Lb, Lactobacillus; Leuc, Leuconostoc; P, Pediococcus; O, Oenococcus

G G 22

The features of this ecosystem will include: (i) physical and chemical properties of the berry surface; (ii) the availability of nutrients for bacterial growth; and (iii) extrinsic influences of the vineyard environmental and management factors including climate, insects and birds, irrigation and application of agrichemicals.

2.4.1 The physical and chemical properties of grape surfaces

Anatomically, the grape berry consists of : (i) the exocarp which is the outermost dermal or skin layer; (ii) the mesocarp which is commonly known as the flesh or pulp; and (iii) the endocarp which represents the innermost part of the mesocarp pulp (near the seeds)

(Hardie et al. 1996). With respect to microbial colonization of the surface, the exocarp is the most relevant component and the following discussion will focus on this layer.

The skin of the grapes forms a heterogenous region constituted by the cuticle, the epidermis and the hypodermis (Fig. 2.3) (Ribéreau-Gayon et al. 2000). Essentially, the cuticle serves as a protective barrier against a variety of stresses (Rosenquist and

Morrison 1989). The cuticle consists of cutin layer and waxes which are embedded within the cuticle (intracuticular wax) or deposited on the outer surface of the cuticle

(epicuticular wax). Cutin, the main structural component of the cuticle, is a lipidic polyester whose principal constituents are 10, 16-dihydroxy hexadecanoic acid and

9,10,18-trihydroxy octa-decanoic acid. Cuticular waxes (intra- and epicuticular wax) are mainly formed by oleanolic acid (C30H48O3), a cyclic tri-terpenoid acid, constituting about 60-80% of the total wax (Commenil et al. 1997). The remaining components are hydrocarbons of chain lengths C14-C32, free alcohols (C20-C34) and free acids (C14-C32), as well as their and aldehydes (Radler and Horn 1965; Commenil et al. 1997;

Casado and Heredia 1999). Some original data on the chemical composition and

G 23

Cuticle Epidermis Exocarp Hypodermis (skin)

Polygonal cells

Pulp

Radial elongated cells Tangential elongated cells Seed

Fig. 2.3 Structure of grape berry at maturity (Ribereau-Gayon et al. 2000) physical organization of cuticle layers can be found in the studies of Chambers and

Possingham (1963), Radler and Horn (1965), Baker (1970), Grncarevic and Radler

(1971), Considine and Knox (1979), Rajaei (1987), Rosenquist and Morrison (1989),

Hardie et al. (1996), Jeffree (1996), Commenil et al. (1997) and Casado and Heredia

(1999).

The physical, chemical and physiological properties of the cuticle have been studied to explain its protective roles (Rosenquist and Morrison 1989; Percival et al. 1993). The epicuticular wax shields the underlying tissue from cuticular transpiration (Schoneherr

1982), attack by microorganisms and insects (Commenil et al. 1997) and injuries arising from wine abrasion, frost and radiation. Also, it affects wettability of the grape surface with water, retention of agrichemical sprays, and adhesive ability of microorganisms.

These functions are related to its hydrophobic nature (Tewari and Skoropad 1976), composition (Marois et al. 1985) and the wax content per unit surface (Rosenquist and

Morrison 1989). G 24

The epicuticular wax consists of an array of overlapping wax platelets (about 0.1 Pm diameter) that form a hydrophobic film or layer, about several microns thick, over the cuticle. The amount, structure, thickness and density of the epicuticular wax layer varies depending on grape cultivar, maturity and environmental factors (Hill et al. 1981; Kök and ÇeĒk 2004). For example, Cabernet Franc grapes have a relatively larger and distinct platelet compared with Riesling grapes, Cabernet Sauvignon berries have more than twice as much cuticle per unit surface area as , , Carignane and

Pinot Noir grapes. Wax deposition is increased in high light, and at higher temperatures and humidity (Rosenquist and Morrison 1989; Percival et al. 1993; Commenil et al.

1997). The epicuticular wax is usually present from the early stages of berry development after flowering. Initially, it appears as a folded layer with distinct ridges, but evolves into a continuous smooth layer of decreased thickness at berry maturity

(Rajaei 1987; Rosenquist and Morrison 1989; Percival et al. 1993; Commenil et al.

1997).

The first contact between bacteria and grape berries normally occurs at this layer of epicuticular wax. This wax layer may interfere with or encourage bacterial colonization and, also it could change the bacterial ecology by limiting diffusion of nutrients and inhibiting the wetting of the grape surface.

2.4.2 Nutrients for bacterial growth on grape surfaces

The chemical composition of the waxy-cuticle surface has already been mentioned. It is not known if these lipidic, hydrocarbon components can serve as growth substrates for microorganisms that are found on the grape surface. However, leaches or exudates of sugars and amino acids from the inner grape tissue (endocarp) to the surface could be a

G G 25 source of nutrients for most microorganisms. Scanning electron microscopy has revealed that most microorganisms, including yeasts and lactic acid bacteria, preferably colonize zones around the peristomatic fractures, as they are able to grow on the small amount of exuded nutrient (Ribéreau-Gayon et al. 2000; Fleet et al. 2002).

Table 2.3 shows the main components of grape tissue and how they change during the maturation of grapes. Immature berries contain very little sugars (2.0 mg/g) such as glucose and fructose, and high contents of malic and tartaric acids (20-30 mg/g), giving the internal tissue a pH of 2.5-3.0. As the berry matures, the total sugar concentration increases to 150-300 mg/g, the concentrations of malic and tartaric acid decrease to 5-

10 mg/g, and the internal pH increases to 3.0-3.5 (Ribéreau-Gayon et al. 2000).

Table 2.3 Main components of tissue of wine grapes

Berry tissue composition Stages of grape Sugars Acidity berry Glucose Fructose Malic acid Tartaric acid development Brix pH TAa (%) q (mg/g) (mg/g) (mg/g) (mg/g) Berry formation 4-6 2-4 0-2 2.5-3.0 1.99-2.75 20-30 15-30 (weeks 1-7) Veraison 8-14 60-70 55-65 3.0-3.3 1.29-1.69 5-10 6-10 (weeks 7-8) Ripening stages 18-22 105-120 115-120 3.3-3.5 0.50-0.71 5-6 5-6 (weeks 9-12) a titratable acidity Data abstracted from several references (Esteban et al. 1999; Barnavon et al. 2000; Varandas et al. 2004)

In unripe grape berries, more than half of the total nitrogen is represented by the cation. As grape matures, especially from veraison, the ammonium concentration decreases, whereas the organic nitrogen increases. Table 2.4 shows the total nitrogen and its relative distribution per berry in the ripening period (Peynaud and

Maurie 1953; Boulton et al. 1995). The total nitrogen in the seeds and skins is relatively high compared with the pulp. Grape juice at maturity contains complex forms of

G G 26 nitrogen compounds (Table 2.4).

Table 2.4 Distribution of total nitrogen in ripe berries and ripe juice of four varieties

Ripe berry Ripe juice % of total N % of total N Variety Total Na Total Nb Poly- Seeds Skins Pulp NH Amino N 4 peptides Cabernet 977 42 43 16 257 19 31 37 13 Sauvignon Merlot 981 49 33 18 285 28 29 28 15 Sauvignon 1291 25 54 22 460 17 39 37 8 Blanc Semillon 849 26 51 23 282 24 39 19 18 a, mg/kg fresh weight; b, mg/L; Data adopted from Boulton et al. (1995)

Several authors have detected sugars, organic acids and amino acids in rinses of the surfaces of healthy grape berries and have suggested that these nutrients originate as exudates from the inner tissue. Kosuge and Hewitt (1964) reported a 10 fold increase in the concentration of reducing sugars in surface rinses of wine grapes as they progressed toward maturity. They also reported amino acids in the rinses, but their total concentration remained relatively constant throughout the maturation process. They suggested that these exudates serve as nutrients for development of the spoilage fungus,

B. cinerea. Padgett and Morrison (1990) conducted a similar study, showing a 10 fold increase in grape surface sugars during maturation. They also reported the high concentration of malic acid and phenolic compounds on the surfaces of immature grapes that decreased substantially during maturation, thereby decreasing their inhibitory influences on the growth of spoilage fungi, such as B. cinerea. Interestingly, they could not detect tartaric acid on the surfaces of grapes, despite its high concentration in inner tissue. They speculated that the process of exudation was compound specific and not an uncontrolled leakage of inner constituents. More recent studies by Varandas et al. (2004) demonstrated the presence of glucose and fructose on

G G 27 grape surfaces, and their increase in concentration during berry ripening. Total sugar content on grape skin represents, on average, 10 r 5% of the equivalent content in juice; however, the concentration of sugars associated with skin varied with the grape variety

(Varandas et al. 2004).

The hydrophobic nature of the grape surface induces water droplet formation.

Consequently, localised areas of exudates with high sugar content and low water activity might occur and have a selective effect on the microflora which grow

(Rousseau and Donèche 2001).

It is evident from the few studies conducted that some constituents of the inner grape tissue find their way to the grape surfaces by general diffusion and leakage through pores, fissures or stomata. These surface constituents may either serve as nutrients that support the growth of microorganisms or they may be inhibitory to microbial growth.

Their influence could possibly be selective, leading to the evolution of different species during grape berry development from immaturity to maturity.

2.4.3 Vineyard environmental factors

The surface of wine grapes could be considered as a hostile habitat for bacteria. The berry surface is exposed to various environmental stresses including rapidly fluctuating temperature and solar radiation, repeated alternation between the presence and absence of free moisture due to rain and dew, and the stresses of agrichemical applications

(Lindow and Brandl 2003). These environmental factors are likely to influence the bacterial ecology of the berry surface (Castelli 1954; Minarik and Ragala 1975). The extremes of temperature, radiation and water availability vary on a large scale with geographical location (latitude, altitude, topography), on a smaller scale within the

G G 28 grape vine canopy, and on a microscopic scale over the grape berry surface (Jackson and Lombard 1993).

The temperature of the grape surface can vary by as much as 15-20qC in one day, and reach values as high as 40-45qC during the period in Australia (Rankine 1989).

Furthermore, different temperature zones can occur across regions and also even with a cluster of grapes, as grapes that are shielded from the sun will be cooler than those directly exposed to sunlight. Hence, these temperature fluctuations can have the effect of imposing a selective force for the more adaptive microorganisms.

The daily exposure to solar UV radiation, including UV-A and UV-B wavelengths, is one of the most prominent features of phyllospheric habitats to which microorganisms must adapt (Beattie and Lindow 1995; Lindow and Brandl 2003). Many epiphytic bacteria are pigmented, and these pigments are presumed to confer protection against

UV radiation (Ayers et al. 1996). A detailed examination of the epiphytic bacteria present on peanut plants exposed to high fluxes of UV, revealed that UV radiation sensitive organisms either died, entered viable non-culturable states or were reduced in number, whereas, populations of more UV tolerant, pigmented strains were increased

(Sundin and Jacobs 1999). UV radiation cause direct DNA damage resulting in inhibition of protein and DNA synthesis which is lethal (Muller-Niklas et al. 1995).

Genes that confer UV tolerance and mutagenic DNA repair have been identified in some epiphytic bacteria (Kim and Sundin 2000; Sundin 2002).

Water is frequently limiting on plant surfaces, especially under high temperature and arid conditions (Hirano and Upper 2000). Decrease in water availability inhibits the

G G 29 growth of many species and selects for those that have adapted to tolerate such stresses

(Rousseau and Donèche 2001). Generally, bacterial isolates from grapes were reported to be more sensitive to low water activity values than yeast strains, although species within Acinetobacter, Brevibacterium and Bacillus showed a wide heterogeneity with respect to water availability (Rousseau and Donèche 2001). The growth of epiphytic bacteria may actively occur following rain, periods of dew and conditions of high humidity (Campbell 1989). High rainfall favours the proliferation of microorganisms on grape berries, particularly oxidative microbiota, with detrimental effects to wine quality

(Longo et al. 1991; Jackson and Lombard 1993; de la Torre et al. 1998).

While each environmental factor has its own impact on bacterial survival and growth, the surface of the grape is simultaneously exposed to a multiple of stresses that are likely to have additive or synergistic consequences on the microbiota (Fleet 1999).

2.4.4 Agrichemical influences

A diversity of agrichemicals is routinely applied throughout grape cultivation to control attack and infection by insects and fungi (Emmett et al. 1992, Cabras and Angioni

2000). These applications are systematically managed from about the time of bud burst until about two weeks before grape harvest, to maximize their effectiveness and minimize carry-over of residues into the juice and wine. Generally, these sprays consist of either chemical or biological agents active against particular insects or fungi, a wetting agent, and water. These agents may influence the structure of the microbial community and physiological development of the grapes, and may be retained as residues on the surfaces of the grapes (Cabras and Angioni 2000). It has been reported that application of agrichemicals could influence the yeast flora associated with grapes,

G G 30 including the incidence of S. cerevisiae (Regueiro et al. 1993; Guerra et al. 1999; van der Westhuizen et al. 2000a, 2000b). Moreover, some of these pesticides (e.g. Euparene,

Mycodifol, Mikal, Cuprosan, Sygan, Acylon Blue) can retard the growth of yeasts and result in stuck or sluggish alcoholic fermentations (Sapis-Domercq 1980; Regueiro et al.

1993). Although the effects of agrichemicals on the bacterial ecology of grapes have not been extensively studied, there is a report that some pesticides (e.g. , dichlofluanid) can hinder the onset of malolactic fermentation by O. oeni (Vidal et al.

2001; Ruediger et al. 2005). Wetting or surfactant agents within these preparations may also influence the microbial community on the surface of grapes. Wetting agents could affect the rates of permeation of agrichemicals through the cuticle by altering the structure of the wax. If sprayed often enough, and in certain combinations, they may cause permanent, acute damage to the delicate waxy surface of the berry, facilitating infection by the fungus, Botrytis cinerea (Marois et al. 1985). Once hyphal elements penetrate berries, subsequent growth brings about further rupture of the epidermal tissues, thereby making the grapes susceptible to secondary infection with bacterial and yeast growth (Fugelsang 1997; Dorado et al. 2001).

Bacillus thuringiensis is a biological insecticide now used extensively in agricultural practices, including application to vines of wine grapes (Emmett et al. 1992; Nester et al. 2002). In Australian vineyards, B. thuringiensis has long been applied as a spray, and gives good to high effective control of attacking pests (Hamblin et al. 1998). It is reported that commercial preparations of B. thuringiensis usually contain a mixture of spores, endotoxin crystals, vegetative cells, cell debris and some carry over of material from the fermentation process used for their production (Glare and O’Callaghan 2000).

There is an abundance of spores and viable vegetative cells in these preparations, and

G G 31 they will be introduced to the surface of wine grapes through application. Introduction of high populations of B. thuringiensis on grapes might lead to competition for nutrients and space, and affect the balance of indigenous microbial flora.

Although firm conclusions cannot be advanced at this stage, it is most likely that agrichemical applications will affect the quantum and diversity of bacterial species associated with wine grapes. The concentration and frequency of application could be important variables in affecting the structure of this bacterial community. The future use of agrichemicals will need to be assessed in terms of their impact on this natural, ecological balance.

2.4.5 Sources of bacterial contamination of grapes

In the vineyard, grapes are exposed to bacteria that occur in the soil, air, wind-blown dust, vegetation, rain, irrigation water, fertilizers and other agrichemical sprays, and bacteria that occur on other parts of the vine such as leaves. Soil serves as a natural or temporary reservoir for a wide range of microorganisms, and wind-blown naturally infested soil is likely to be one of the main sources of microorganisms on the surface of grape berries. Recently, Whitelaw-Weckert et al. (2004) reported the presence of species of Bacillus and Pseudomonas and cellulolytic bacteria in soil from an Australian vineyard. Although soils are not considered a major source of lactic acid bacteria, species of Enterococcus (E. avium, E. mundtii, E. facium, E. hirae), Lactococcus lactis and sporeforming lactic acid bacteria (e.g. Sporolactobacillus inulinus) have been isolated from the soil of rhizospheres of vines in Taiwan and Japan by enrichment culture (Chen et al. 2005).

G G 32

There is strong view that insects are probably one of the most significant sources of bacterial contamination of grapes, although specific details of this relationship require definition (Mortimer and Polsinelli 1999; Fleet 2003b). Many insects including honey bees, halictus bees, syrphid flies (Scaeva pyrastri) and wasps visit grape blossoms. They are attracted to the flowers, primarily for pollen and nectar, and sometimes gather honeydew from grape vine leaves. Various species of acetic and lactic acid bacteria

(Bifidobacterium, Lactococcus, Lactobacillus, Leuconostoc) can be isolated from the bodies and intestinal contents of bees and wasps (Ruiz-Argueeso and Rodriguez-

Navarro 1973; Gilliam 1997; Reeson et al. 2003). Other bacterial species associated with honey bees were reported by Gilliam (1997). Gram-variable pleomorphic bacteria,

Bacillus spp. and Enterobacteriaceae were consistently found, although the intestinal microflora of mature, worker bees may vary somewhat with the age of the bee, season, and geographical location (Gilliam and Valentine 1974). In addition to the insects just mentioned, there is a diversity of ants, moths, mites, mealybugs, weevils, cicadas, locusts, spiders and grasshoppers that visit, infect and attack grape vines and grape berries at various stages of their development (Buchanan and Amos 1992). It is not uncommon to see these insects crawling over bunches of berries as they grow and mature on the vine. It is most likely that these visitors will contribute generally and specifically to the bacterial flora of grapes, but no information is available on this topic.

Birds also attack grape berries, especially as they approach maturity (Boudreau 1972;

Buchanan and Amos 1992). They will contribute a diversity of bacterial contamination but, most significantly, they damage the waxy-cuticle layers, allowing access to nutrients for their growth.

Although commercial concentrates of agrichemicals are generally sterile or have low

G G 33 microbial loads, they need to be reconstituted in water for spray application. Usually, this is farm water (dam, river, bore) that is not sterile. Moreover, reconstituted preparations may be used over a period of several hours or more, allowing time for microbial contaminants to grow. Ng et al. (2005) demonstrated rapid and significant bacterial growth in a range of agrichemical preparations after they were reconstituted in supplies of farm water. Apart from the active ingredient, agrichemical preparations usually carry other adjunct agents to enhance their functionality, and these could serve as a source of nutrients for bacterial growth. Thus, agrichemicals could be an unrecognized source of microbial contaminants and nutrients on grapes.

2.4.6 Bacterial properties

As mentioned already, the grape surface presents an environment of unique chemical and physical attributes – essentially a waxy, hydrophobic surface that is subject to significant diurnal and seasonal fluctuations in temperature, sunlight, irradiation and water availability. To successfully colonize this habitat, bacteria will need to express some equally unique survival and growth mechanisms. Multiple traits that influence the growth and survival of bacteria on phyllospheric habitats include motility, the ability to withstand physical removal from the plant surface through the action of rain, dew deposition, irrigation water and wind, and protection mechanisms against environmental factors such as temperature, UV radiation, and desiccation. The adhesion, attachment, and aggregation of microbial cells to phyllospheric locations have been correlated with their production of surface structures or molecules (Romantschuk 1992). These structures and molecules mainly include (i) filamentous proteinaceous appendages called fimbriae (pili) and fibrillar molecules composed of cellulose (Korhonen et al.

1986), and (ii) other extracellular polymeric secretions (EPS) (Romantschuk 1992;

G G 34

Beattie 2002). Fimbriae (pili) and fibrillar materials are thought to be associated with primary attachment, interacting with receptors, which are the glycoproteins or on the leaf or fruit surface. In addition, fibrils of bacterial cellulose assist cluster formation, and cell aggregation (Romantschuk 1992; Davey and O’Toole 2000). Species of Gram- negative bacteria such as Pseudomonas, Erwinia, Pantoea (Enterobacter), and

Xanthomonas are well known for these properties and this might explain their frequent occurrence at phyllospheric habitats (Romanschuk 1992). Many epiphytic bacteria, including plant pathogens, are known to produce a diverse array of EPS that enhance their adhesion to solid surfaces (Gross and Rudolph 1987; Denny 1995). EPS have a multitude of binding and buffering properties that could lead to the establishment of a micro-niche on leaf, fruit and vegetable surfaces and may also function in quorum sensing responses to control bacterial population densities (O’Toole et al. 2000;

Morris et al. 2002; Morris and Monier 2003). In addition to enhancing attachment and proliferation of cells at the phyllosphere, EPS also protect the organisms from desiccation (Ophir and Gutnick 1994) and assist the chelation of heavy metals

(Whitfield 1988). Schönherr and Baur (1996) have suggested that EPS can absorb and concentrate volatile compounds exuded from aerial plants, and this could be a selective factor on the species which grow (Thompson et al. 1993; Jurkevitch and Shapira 2000).

Bacterial species frequently found on the phyllosphere are usually chromogenic and, produce pigments that are believed to protect the cells against sun light and UV radiation as mentioned previously. Organisms producing orange to pink and red carotenoid pigments have significantly higher levels of UV tolerance compared to non- pigmented isolates (Sundin and Jacobs 1999). Epiphytic bacteria can also produce numerous compounds that enhance plant cell leakiness, and facilitate a supply of nutrients (Morris et al. 2002). For example, production of auxin, toxins such as

G G 35 syringomycin and syringopeptin, pectolytic , and biosurfactants by species of

Pseudomonas lead to greater availability of nutrients from plant cells (Durbin 1983;

Alghisi and Favaron 1995; Glickman et al. 1998).

Bacteria respond to environmental stresses by activating survival mechanisms (Roszak and Colwell 1987; Morita 1993; Barcina et al. 1997). While some can survive by forming resistant spores, nonsporulating microorganisms are able to persist by entering a viable but nonculturable (VBNC) state in response to extremes of temperature, pH, ultraviolet light, oxygen concentration, water availability, solute concentration and nutrient limitation (Oliver 1993, 2000; Colwell 2000). The VBNC state is defined as cells which are metabolically active but unable to undergo the cellular division for growth in liquid or agar medium (Oliver 1993, 2000; Stephens 2005). The surface of grapes represents a unique phyllospheric habitat for microorganisms and, as mentioned before, is subject to repeated, and rapid alteration of adverse environmental conditions.

Therefore, it is likely that organisms in a VBNC state will occur on grape surfaces. The putative VBNC state has been reported in many genera, mainly in Gram-negative organisms, but also including Gram-positive species such as Micrococcus luteus,

Listeria monocytogenes, and Enterococcus faecalis (Millet and Lonvaud-Funel 2000;

Stephens 2005).

2.5 BACTERIA ASSOCIATED WITH WINE GRAPES

The significance and roles of fungi, yeasts and bacteria in wine production have already been mentioned in Section 2.3. The following section will focus on the bacteria associated with wine production, in particular those studies that have considered the

G G 36 bacterial species found on wine grapes. As mentioned before, species of lactic acid bacteria and acetic acid bacteria are most important in winemaking but, on occasions, other species can be significant.

2.5.1. Lactic acid bacteria

Lactic acid bacteria have three impacts on wine production. They conduct the MLF, they can cause wine spoilage and some species can cause stuck or sluggish alcoholic fermentations (Fleet 2001, 2003a). Their significance in wine production has been known since the early observations of Louis Pasteur about 150 years, and they have been extensively studied since that time. This literature has been thoroughly reviewed and the reader is referred to Kunkee (1967, 1974), Amerine and Kunkee (1968), Lafon-

Lafourcade and Ribéreau-Gayon (1984), Wibowo et al. (1985), Henick-Kling (1993,

1995), Liu and Pilone (1998), Lonvaud-Funel (1999), Fleet (2001), Liu (2002), and

Bartowsky (2005).

2.5.1.1 Isolation from wines

The vast majority of studies of lactic acid bacteria have been concerned with isolates made at various stages during fermentation and storage. Table 2.5 lists the species of lactic acid bacteria that have been isolated from wines undergoing MLF, or spoilage.

Three genera of lactic acid bacteria were originally considered to occur in wine;

Leuconostoc, Lactobacillus and Pediococcus. Gram positive cocci or cocci- belonging to the genus Leuconostoc were frequently associated with wine as part of the normal bacteriological flora and as participants in the malolactic fermentation (Bidan

1956; Radler 1958, Luthi and Vetsch 1959; Ingraham et al. 1960; Peynaud and

Domercq 1968), but none of theses reports satisfactorily identified them as existing

G G 37

Table 2.5 Species of lactic acid bacteria isolated from wines

Species of lactic acid bacteria References Lactobacillus spp. 33, 28, 31, 4, 16, 6 Lb. brevis 17, 19, 9, 26, 32, 3, 23, 4, 10, 29, 13, 8, 7 Lb. buchneri 9, 23, 4, 6 Lb. casei 32, 23, 6, 2, Lb. cellobiosis 4, 6 Lb. collinoides 5 Lb. delbrueckii 19, 22 Lb. desidiosus 26 Lb. dextranicum 3 Lb. fermentum 23 Lb. fructivorans 25, 24, 5, 13 Lb. hilgardii 17, 19, 9, 26, 23, 4, 22, 8, 24, 5, 13, 7 Lb. jensenii 4 Lb. kunkeei 14 Lb. leichmanii 9 Lb. mali 5 Lb. nagelii 15 Lb. paracasei 7 Lb. plantarum 33, 3, 23, 6, 29, 13, 30 Lb. vermiforme 30, 7 Lb. zeae 30 Leuc. citrovorum 27 Âre-confirmed as Leuc. oenos by Garvie (1967) Leuc. citrovorum 21 Leuc. gracile 2, 25 Leuc. mesenteroides 18 Âre-confirmed as Leuc. oenos by Garvie (1967) Leuc. mesenteroides 16, 21 Leuc. oenos/O. oeni 25, 32, 1, 3, 23, 20, 16, 6, 10, 22, 29, 12, 24 Leuconostoc spp. 28, 31, 2. 19 P. cerevisiae 9, 32, 23, 4, 16 P. curvatus 29 P. inopinatus 11 P. parvulus 6, 11 P. pentosaceus 32, 23, 4, 22 Pediococcus spp. 19, 28, 31 1, Beelman et al. (1977); 2, Bidan (1956); 3, Chalfan et al. (1977); 4, Costello et al. (1983); 5, Couto and Hogg (1994); 6, Davis et al. (1986); 7, du Plessis et al. (2004); 8, Dicks and van Vuuren (1988); 9, du Plessis and Van Zyl (1963); 10, Edinger and Splittstoesser (1986); 11, Edwards and Jensen (1992); 12, Edwards et al. (1991); 13, Edwards et al. (1993); 14, Edwards et al. (1998b); 15, Edwards et al. (2000); 16, Fleet et al. (1984); 17, Fornachon (1957); 18, Fornachon (1964); 19, Ingraham et al. (1960); 20, Lafon-Lafourcade et al. (1983); 21, Luthi and Vetsch (1959); 22, Manca de Nadra and Strasser de Saad (1987); 23, Pan et al. (1982); 24, Pardo and ZuĔiga (1992); 25, Peynaud and Domercq (1968); 26, Peynaud and Domercq (1970); 27, Radler (1958); 28, Rankine (1977); 29, Sieiro et al. (1990); 30, Stratiotis and Dicks (2002); 31, van der Westhuizen et al. (1981); 32, Weiller and Radler (1970); 33, Yoshizumi (1963)

G G 38 species with in Leuconostoc. Later, they were considered to form a new species which was named Leuconostoc oenos by Garvie (1967), but subsequently this organism was reclassified as Oenococcus oeni by Dicks et al. (1995). Principally, O. oeni and possibly

Lb. plantarum are the main species that conduct a MLF which usually enhances wine quality through deacidification, amelioration of flavour and increase in microbiological stability (Bartowsky 2005). Other species of lactic acid bacteria usually have undesirable consequences, and contribute to problems of spoilage, taints and stuck/sluggish fermentations (Wibowo et al. 1985, Sponholz 1993; Bisson 1999;

Edwards et al. 1998a, 1999; Lonvaud-Funel 1999; du Toit and Pretorius 2000).

2.5.1.2 Isolation from grapes and musts

It is generally thought that the lactic acid bacteria of wine production originate from the grapes. Table 2.6 summarises the species of lactic acid bacteria that have been isolated directly from aseptically analysed grapes or the musts produced from freshly harvested grapes. The musts were not necessarily processed under aseptic conditions, so there is a possibility that the lactic acid bacteria reported come from sources other than the grape surface. The early literature certainly shows that lactic acid bacteria can be isolated from wine grapes. Kunkee (1967) cites work by Nessler who detected the presence of these bacteria on grape skins. Rankine and Bridson (1971) reported that French researchers in the Gironde region of Bordeaux, found lactic acid bacteria in 16-25% of samples of ripe grapes taken aseptically from vineyards. Similarly, Ribéreau-Gayon and

Peynaud (1975) indicated that grape skins, including the grape pellicles were a predominant source of wine microflora, although the malolactic bacteria were less widespread on vineyard grapes than yeasts and acetic acid bacteria. Kunkee et al.

(1965) reported that spontaneous malolactic fermentation occurred in wine produced

G G 39 under “semi-sterile” laboratory conditions, strongly suggesting a grape source of the

MLF. However, there are contrasting reports where MLF did not occur in wines prepared from aseptically managed grapes, suggesting that grapes may not be a consistent or predictable source of these bacteria (Webb and Ingraham 1960; Peynaud and Domercq 1961).

These earlier findings are more or less confirmed in more recent studies. Lafon-

Lafourcade et al. (1983), Manca de Nadra and Strasser de Saad (1987), Sieiro et al.

(1990) and de la Torre et al. (1998), all experienced difficulty in detecting lactic acid bacteria on wine grapes. Samples were either negative for the presence of these bacteria or populations were very low (less than 50 CFU/mL of grape homogenate). Various species of Lactobacillus were isolated on these occasions (Table 2.6). Suárez et al.

(1994) examined the populations and species of lactic acid bacteria on several grape varieties as they matured through veraison. Low populations (102-104 CFU/g) were generally reported, with Lb. plantarum and Leuc. dextranicum being the main species.

The populations increased as the grapes approached harvest maturity, and for Cabernet

Sauvignon, counts at 106 CFU/mL were reported. However, this study did not give clear descriptions of its sampling and analytical procedures. Weiller and Radler (1970) reported the isolation of lactic acid bacteria such as Lb. plantarum and Leuc. mesenteroides from leaves of grapevines, but not from fresh grape juice. There appears to be no definitive report on the isolation of the malolactic bacterium, O. oeni (Leuc. oenos) from the surface of grapes.

There is no difficulty in detecting lactic acid bacteria in freshly processed grape musts

(Table 2.6). Populations of 102-104 CFU/mL are usually reported. Moreover, a great

G G 40 diversity of species is found, including the presence of O. oeni. Presumably, the processing operations used to crush the grapes add to their microbiota.

Table 2.6 Species of lactic acid bacteria isolated from grapes and musts

Samples Species of lactic acid bacteria Reference Grapes Lb. brevis 11 Lb. buchneri 10 Lb. casei 5, 11 Lb. curvatus 10 Lb. hilgardii 9, 5, 11 Lb. plantarum 12, 5, 3, 11 Leuc. dextranicum 11 Leuc. mesenteroides 12 Musts Lb. brevis 9, 5, 8 Lb. casei 5 Lb. confuses 8 Lb. hilgardii 8, 9 Lb. plantarum 1, 2, 5, 8 Leuc. mesenteroides 4, 5 Leuc. oenos 1, 5 Leuc. paramesenteroides 8 P. cerevisiae 4 P. pentosaceus 7 1, Costello et al. (1983); 2, du Plessis et al. (2004); 3, Edinger and Splittstoesser (1986); 4, Fleet et al. (1984); 5, Lafon-Lafourcade et al. (1983); 6, Lonvaud-Funel and Strasser de Saad (1982); 7, Manca de Nadra and Strasser de Saad (1987); 8, Pardo and ZuĔiga (1992); 9, Peynaud and Domercq (1970); 10, Sieiro et al. (1990); 11, Suárez et al. (1994); 12, Weiller and Radler (1970)

2.5.1.3 Isolation from winery environments

It is well recognized that the winery environment (e.g. surfaces of winery equipment, pump, hoses, walls, barrels) is a significant source of indigenous yeasts in wine production (Fleet et al. 2002). A similar conclusion could be drawn about the lactic acid bacteria of wine production (Kunkee 1967, 1974). As mentioned already, there is little difficulty in isolating lactic acid bacteria from winery produced grape juice or must, and spontaneous development of the MLF in wines due to the growth of O. oeni is a common occurrence. Like S. cerevisiae, O. oeni could take up residence in the winery environment, its presence being selected by its tolerance to the more stressful properties

G G 41 of wine and grape juice. Presumably, the initial inoculation for this organism would come from the grapes or vineyard ecosystem.

Very few studies have been conducted on the winery environment as a source of lactic acid bacteria. Earlier studies reported malolactic bacteria in the winery environment such as floors, walls and wooden barrels (Webb and Ingraham 1960;

Ribéreau-Gayon and Peynaud 1975). Lactobacillus sp. have been isolated from winery bottling equipments and the bottling room atmosphere (Donnelly 1977). Although this finding is not comprehensive or conclusive, species of Enterococcus (E. avium, E. mundtii, E. faecium, E. hirae) and Lc. lactis have been isolated from the soil of rhizospheres of vines in Taiwan and Japan (Chen et al. 2005).

2.5.2 Acetic acid bacteria

The acetic acid bacteria, Acetobacter and Gluconobacter are linked by their common ability to cause vinegary spoilage of grapes, musts and wines. Despite their significance, there have been few ecological studies of their occurrence on grapes. Sound, unspoiled grapes harbour acetic acid bacteria at low populations, generally at about 101-103 CFU/g, with Gluconobacter oxydans being the predominant species at this stage (Joyeux et al.

1984; Drysdale and Fleet 1988; Sponholz 1993; du Toit and Pretorius 2000). Damaged, spoiled grapes and those infected with the mould, B. cinerea, harbour much higher populations (approx. 106 CFU/g) that are characterized by a dominance of the species of

A. aceti and A. pasteurianus (Lafon-Lafourcade and Ribéreau-Gayon 1984; Joyeux et al.

1984). However, other studies have determined the predominance of G. oxydans or both

G. oxydans and A. aceti on spoiled and Botrytis-infected grapes (Barbe et al. 2001; du

Toit and Lambrechts 2002; González et al. 2005).

G G 42

The population of acetic acid bacteria in freshly extracted must reflects those populations present on the grapes (Joyeux et al. 1984; Lafon-Lafourcade and Ribéreau-

Gayon 1984). The studies of Joyeux et al. (1984) suggest that acetic acid bacteria show little tendency to grow in grape juice during alcoholic fermentation and undergo a substantial reduction in population. At the end of alcoholic fermentation, their population is generally less than 102-103CFU/mL. However, in some musts having a high pH (>3.7), the population of acetic acid bacteria was reported to be increase to 104-

105 CFU/mL during fermentation (du Toit and Lambrechts 2002). Drysdale and Fleet

(1989a, 1989b) also reported the potential for acetic acid bacteria to grow in grape juice/must and cause stuck alcoholic fermentation.

G. oxydans is rarely isolated from wines at this stage, and A. aceti, A. pasteurianus, A. liquefaciens and A. hansenii are the main species found (Joyeux et al. 1984; Drysdale and Fleet 1988; du Toit and Lambrechts 2002; Bartowsky et al. 2003). Subsequent transfer of the wine from fermentation tanks to other storage vessels may produce sufficient agitation and aeration to encourage the growth of surviving species of acetic acid bacteria (Joyeux et al. 1984). The distribution of Acetobacter spp. in wines can be related to the country of origin (du Toit and Pretorius 2000). In Australian wines, A. pasteurianus was most frequently isolated, whereas in France and the USA, A. aceti was dominant (Vaughn 1955; Drysdale and Fleet 1985, 1988). The occurrence and isolation of acetic acid bacteria from wine environments are summarized in Table 2.7.

G G 43

Table 2.7 Acetic acid bacteria isolated from grapes, musts and winery environments

Samples Bacterial species Reference Grapes A. aceti, A. paradoxum, A. ascendens, A. 4, 6, 8, 9, 10, 11, 12 rancens, A. mesoxydans, A. pasteurianus, A. suboxydans, A. xylinum, G. oxydans, Gluconoacetobacter hansenii Musts G.. oxydans, A.. aceti, 5, 8, 10 A.. pasteurianus, A. liquefaciens. Wine A. pasteurianus, A. aceti, 1, 2, 3, 5, 6, 7, 8, 11, 12 A. hansenii, Gluconobacter sp. G. oxydans 1, Bartowsky et al. (2003); 2, Barbe et al. (2001); 3, Drysdale and Fleet (1985); 4, Drysdale and Fleet (1988); 5, du Toit and Lambrechts (2002); 6, du Toit and Pretorius (2000); 7, González et al. (2005); 8, Joyeux et al. (1984); 9, Lafon-Lafourcade and Joyeux (1981); 10, Lafon-Lafourcade and Ribéreau-Gayon (1984); 11, Sponholz (1993); 12, Vaughn (1955)

2.5.3 Other bacteria

Other bacteria reported to be significant in wine production include species of Bacillus,

Clostridium, Actinomyces, and Streptomyces (Gini and Vaughn 1962; Murrell and

Rankine 1979; Sponholz 1993; Fleet 1998, 2001). Their presence is associated with spoilage and the production of off-flavour, but such occurrences are quite rare. Grapes could be a source of such species, but very little investigations have addressed the incidence of bacteria on grapes other than lactic acid and acetic acid bacteria. In line with other phyllospheric habitats (Billing 1976), it would not be unexpected to find a greater diversity of bacterial species on wine grapes. Bell et al. (1995) noted the isolation of endophytic bacteria from the sap of grapevines. Seventy eight percent of the isolates were Gram negative bacteria, with Pseudomonas and Enterobacter being the predominant genera. After these two genera, Rahnella aquatilis, Pantoea agglomerans,

Staphylococcus spp., Rhodococcus luteus and Curtobacterium flaccumfaciens were the most frequently isolated bacterial species. It would not be unreasonable to expect such species to contaminate the surface of grapes. Other bacterial species (Acinetobacter,

Brevibacterium, Bacillus) isolated from healthy and rotten grapes have been also mentioned by Rousseau and Donèche (2001). Subden et al. (2003) reported the

G G 44 presence of bacteria in the musts of Riesling grapes as processed for icewine production in Canada. Bacterial populations in the freshly produced must ranged from 104-105

CFU/mL. The following species and their proportion (%) of the total population were reported: Pantoea agglomerans (44%), Curtobacterium flaccumfaciens (29%),

Pseudomonas corrgata (20%) and Curtobacterium pusillum (7%). The absence of acetic acid bacteria and lactic acid bacteria in their isolation was notable.

2.6 METHODS FOR THE ANALYSIS OF BACTERIA ASSOCIATED WITH WINE GRAPES

2.6.1 Conventional microbiological analysis of wine grapes

The standard approach to study the bacteria associated with wine grapes has been to rinse the surfaces of grapes, or homogenize the grapes, and then test for bacteria in the rinses or homogenates by plating onto appropriate agar media. In some cases, grape samples have been incubated in liquid enrichment media prior to plating onto an agar medium in order to detect bacterial species present at very low populations. Common media used for the isolation of wine bacteria have been reviewed (Pan et al. 1982;

Wibowo et al. 1985; Drysdale and Fleet 1988; Fleet 1993; Fugelsang 1997). They include de Man Rogosa Sharpe (MRS) agar, tomato juice agar, tomato juice wine agar,

Irrmann agar, Nakagawa agar, agar and various modification of MRS agar for lactic acid bacteria, and GYC agar, Mannitol agar, WL Nutrient agar with 2% ethanol, and Frauter and Carr agar for acetic acid bacteria. It was reported that no single medium can reliably recover all species of bacteria associated with wines or grapes, therefore, the use of two or more isolation media is often recommended (Fleet 1993).

G G 45

The conventional cultural approach can produce valuable and useful data, but there are many factors that could cause erroneous ecological information. Some of these factors include inhibitory tissue extracts in homogenates or rinses, influence of diluent composition on organism viability, long time span between dilution steps and plating, failure of enrichment cultures to amplify a desired population, and poor anaerobic methodology. Biases and cell death during these procedures may result in poor reproducibility and a distortion of the ecological truth (Fleet 1999). Also, it is now well recognized in the field of microbial ecology that many microorganisms in natural ecosystems may not be culturable on agar isolation media (Oliver 1993). The presence of VBNC microorganisms poses another problem in the use of conventional cultural methods. As mentioned in Section 2.4.6, VBNC cells are those that, after exposure to adverse conditions, enter a phase whereby they cannot produce a colony on usual growth media despite the fact that they are physiologically viable. Thus, such cells cannot be detected using conventional methods (Kurath and Morita 1983). A study by

Millet and Lonvaud-Funel (2000) demonstrated that both wine lactic acid bacteria and acetic acid bacteria enter a VBNC state during wine storage. Although these cells are capable of returning to their normal healthy state upon removal of the responsible environmental stresses, the VBNC state may be the reason why these bacteria spoil wine during storage but, sometimes, are not detectable by conventional techniques. In addition, microorganisms that grow on isolation media are likely to be the ones most adapted for growth under such conditions, and not necessarily the ones that are most metabolically active or abundant in the environment (Embley and Stackebrandt 1996).

This means that current knowledge about the microbial profile of many habitats, obtained using culture-based methods, is likely to be over-simplified and may represent a significant under-estimation of the true ecology.

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To overcome the limitations of culture-based methods for analysing the bacterial ecology of natural habitats, culture-independent, molecular techniques have been developed in recent years (Head et al. 1998; Torsvik et al. 1998; Denton et al. 1999;

Ercolini et al. 2001).

2.6.2 Molecular strategies for monitoring bacterial communities

Molecular methods are now routinely used to identify bacterial isolates to genus and species, and to differentiate strains at the sub species level. Such methods based on 16S rRNA sequence analysis, are now increasingly used in ecological studies (Embley and

Stackebrandt 1996). In addition, new, culture-independent molecular approaches, based on PCR amplification of isolated DNA and analysis of the amplicons by denaturing gradient gel electrophoresis (DGGE) or temperature gradient gel electrophoresis

(TGGE) are now used to investigate the ecological profile of microorganisms in natural habitats (Head et al. 1998; Muyzer and Smalla 1998) including food ecosystems

(Ercolini 2004; Giraffa 2004). The basic strategy is outlined in Fig. 2.4. Total DNA is extracted from samples of the product. Using primers targeting conserved regions, bacterial ribosomal DNA within the extract is specifically amplified with PCR.

Generally, highly variable regions in bacterial 16S rDNA are targeted, and other regions such as the 23S rDNA and 16S-23S rDNA spacer (ITS) may be used. The amplicons produced by PCR are separated by DGGE, isolated and identified by sequencing. The basis of the technique is separation of DNA fragments of similar length, but different in base composition, according to their melting properties in a denaturing gradient gel

(Fischer and Lerman 1983; Myers et al. 1985). Amplified fragments of up to about 500 bases with only one or a few differences in sequence are easily separated from each other (Nübel et al. 1996).

G 47

Food / Beverage/ Natural habitat

Extract and purify total DNA

PCR amplification of bacterial ribosomal DNA

DGGE separation of DNA amplicons

Isolate and sequence bands to give species identity

Fig. 2.4 Flow chart of procedure for profiling and identifying bacteria in habitats by PCR-DGGE

PCR-DGGE can be applied to analyse bacterial communities in complex environments such as foods, beverages, soils, phyllospheres and rhizospheres (Gelsomino et al. 1999;

Heuer and Smalla 1997, 1999; Smalla et al. 2001; Giraffa 2004). The band patterns generated by DGGE analysis of the mixture of PCR-amplified rDNA fragments can provide an initial ecological overview of the most dominant species, as each DNA band found in the gel usually represents one bacterial species in the microbial ecosystem. The band is excised from the gel and reamplified to increase its concentration, and then sequenced to give the species identity. Thus, a profile of the species associated with the ecosystem is obtained without the need for culture. It is believed that this culture- independent molecular method overcomes the biases and limitations of culture methods, and reveals species that might occur as dormant and possibly unculturable forms

(Wintzingerode et al. 1997). Consequently, a more accurate representation of the habitat ecology is obtained (Muyzer and Smalla 1998). Using reverse transcriptase-PCR to target extracted RNA, a profile of metabolically active species, as opposed to the dead G 48 or non-viable flora, can be obtained. PCR-DGGE is being increasingly applied to determine the microbial flora of foods, beverages and natural habitats. In several cases, it could be concluded that PCR-DGGE revealed much previously unknown information about the microbial community in ecosystems (Smit et al. 2001; Yang et al. 2001).

Hence, it can be anticipated that its application to the study of wine grapes offers great potential to increase the understanding of this ecosystem.

2.6.3 Factors affecting the performance of molecular methods for

analysis of bacteria

Despite the potentially new and valuable information that can be gained from molecular methods in microbial ecological studies, there are some limitations that need to be recognized. In regards to the PCR-DGGE approach, errors and bias can occur during nucleic acid extraction, PCR, DGGE and sequence analysis.

2.6.3.1 DNA extraction

In the application of molecular methods to study the organisms in natural environments, one of the first hurdles is the recovery of total, representative microbial DNA. The purity and concentration of template DNA can be critical when it is used in PCR assay.

Ideally, extraction of DNA should produce a high yield of DNA of sufficiently good quality. The microbial community within the habitat can exist in several morphological forms including mycelia, spores, vegetative unicells, injured cells and cysts (Krsek and

Wellington 1999), as well as in different physiological stages (e.g. exponential, stationary, autolysing, dead). These variable morphological and physiological forms of cells at the time of assay could lead to insufficient or preferential disruption of cells, and affect the efficiency of DNA extraction and the quality of the DNA template for PCR.

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Vigorous conditions are usually required for cell lysis of Gram-positive bacteria and, simultaneously, they might contribute to highly fragmented nucleic acids from Gram- negative cells. Fragmented nucleic acids are sources of artifacts in reverse transcription or PCR amplification experiments and may contribute to the formation of chimeric PCR products (Liesack et al. 1991).

Methods for extraction of DNA from environmental samples can be divided into physical (bead-beating, sonication, freezing-thawing), and chemical based methods

(using chemicals such as SDS, phenol, various detergents, or lysozyme, proteinase K, pronase). Practically, a combination of physical and chemical extraction methods, such as lysozyme and detergent lysis treatment with SDS followed by bead-beating was used to achieve high cell lysis efficiency (Krsek and Wellington 1999). Nevertheless, critical evaluation and some degree of standardization of these methods are needed.

2.6.3.2 PCR

Due to its inherent simplicity, speed and sensitivity to small amounts of DNA, PCR has established itself as a powerful molecular tool for studying the ecology of bacteria

(Giovannoni 1991; Birt and Baker 2000). Theoretically, PCR can detect as low as one copy of a target DNA molecule, however, problems of inhibition of PCR amplification for target DNA in total DNA extracts are often encountered when working with food or natural ecosystems. These problems are largely due to plant , humic components and other polyphenolic materials which can co-purify with the target DNA

(Wilson 1997, Marshall et al. 2003). In addition, the relatively low amounts of the desired bacterial species among the high background/non-target flora (e.g. filamentous fungi, yeast) or presence of the host DNA (e.g. plant) are other factors that influence the

G G 50 efficiency of the PCR. Along with inhibition of PCR, selective amplification of DNA extracted from mixed communities may bias the analysis of bacteria in food and natural ecosystems. Selective amplification can be due to differences in size, copy number of target gene and G+C content of the template DNA, accessibility of the templates to primer hybridization after denaturation (secondary structures), efficiency of primer-template hybrid formation (primer preference), and concentration of templates

(Farrelly et al. 1995; Suzuki and Giovannoni 1996; Zheng et al. 1996; Polz and

Cavanaugh 1998; Kanagawa 2003). The formation of chimeric PCR products, or fusion of two different sequences, and the misincorporation of bases by Taq polymerase during

PCR are both possible events (Cariello et al. 1991; Eckert and Kunkel 1991;

Speksnijder et al. 2001; Qiu et al. 2001).

Inhibition and differential amplification, and formation of PCR artefacts can cause misleading and false data and, ultimately, distortion of the ecological truth. Good discussion of these pitfalls can be found in Farrelly et al. (1995), Suzuki and

Giovannoni (1996), Wang and Wang (1997), Wilson (1997), Wintzingerode et al.

(1997), Fogel et al. (1999), Speksnijder et al. (2001) and Kurata et al. (2004).

2.6.3.3 DGGE

A size limitation exists for DGGE analysis in that it is mostly suitable for the separation of DNA fragments fewer than 500 nucleotides. This poses a restriction on the amount of sequence information that can be analysed for phylogenetics and probe design (Myers et al. 1985; Muyzer and Smalla 1998). Ideally, one species yields one band in a DGGE gel, but, two or even more bands have been detected for some strains. This may be due to sequence heterogeneities of 16S rDNA operons (Nübel et al. 1996; Fogel et al. 1999).

G G 51

Also, multiple banding patterns caused by heteroduplex bands, can be obtained during mixed-template PCR when annealing occurs between template DNA with high sequence similarity (Ferris and Ward 1997; Wintzingerode et al. 1997). Several bands from one species can be wrongly interpreted as the presence of species with diversity.

To overcome multiple banding patterns caused by the 16S rDNA heterogeneity, the gene for the RNA polymerase beta subunit, rpoB, is suggested to be the PCR target.

However, the low number of sequences currently available in gene libraries is a limitation for using this gene for species identification (Dahllöf et al. 2000, Dahllöf

2002). Co-migration of DNA fragments with similar melting points can be another problem because single DGGE band might not represent a single microorganism (Heuer and Smalla 1997; Sekiguchi et al. 2001). For ecosystems containing large numbers of equally abundant microorganisms, highly complex banding patterns may be obtained, impeding the qualitative and quantitative analyses by DGGE gels (Wintzingerode et al.

1997).

2.6.3.4 Interpretation of 16S rRNA sequence data

The ultimate goal of a PCR-mediated analysis of 16S rRNA molecules from complex microflora is the retrieval of sequence information, which allows determination of microbial diversity. The quality of results obtained by comparative 16S rRNA sequence analyses strongly depends on the available database. Although more than 75,000 full and partial 16S rRNA sequences are publicly available, those from cultivated microorganisms or uncultured DNA (e.g. DNA band in DGGE) from environmental samples often exhibit a low sequence similarity to known sequences making their phylogenetic affiliation difficult (Wintzingerode et al. 1997; Giraffa and Neviani 2001;

DeSantis et al. 2003). This causes questions as to whether environmental sequences

G G 52 represent uncultured, or novel microorganisms, or whether they cannot be assigned to known taxa due to the fact that, for many cultivated microorganisms, 16S rRNA sequences are not yet available or of low quality (Wintzingerode et al. 1997).

2.7 SUMMARY

Bacteria impact on the quality and efficiency of wine production. Grapes can be a significant source of bacteria in wine production. Most studies to date have focused on the association of lactic acid bacteria and acetic acid bacteria with wine grapes.

However, a variety of other bacterial species may be present at this habitat. The surface of the grapes is a phyllospheric habitat, and the associated bacterial species are likely to evolve and change as the grape berry matures on the vine in response to environmental conditions and influences of viticultural practices. Methods for analysing the bacterial ecology of wine grape have so far been based on the use of standard cultural procedures.

These procedures have well-known limitations that may underestimate the true microbial ecology of natural habitats, such as wine grapes. Such limitations may be overcome by using new culture-independent, molecular methods for ecological analyses.

G G 53 CHAPTER THREE

BACTERIAL ECOLOGY OF WINE GRAPES DURING CULTIVATION AT SEVERAL VINEYARDS IN AUSTRALIA

3.1 INTRODUCTION

The surface of grapes represents a unique phyllospheric habitat for yeasts, bacteria and filamentous fungi that influence the quality and efficiency of wine production (Fleet

2003a). Compared to yeasts (Fleet et al. 2002) and fungi (Emmett et al. 1992;

Fugelsang 1997), little attention has been given to the bacteria associated with grapes.

Grapes represent a primary source of bacteria that enter the winery environment where their impacts on the malolactic fermentation and wine spoilage are well recognised

(Fugelsang 1997; Fleet 1998). In some circumstances, such bacteria can interfere with the growth of yeasts and cause stuck or sluggish alcoholic fermentation (Bisson 1999;

Edwards et al. 1999). As part of the natural process of microbial interactions, bacteria at the grape surface may influence the presence and populations of yeasts and filamentous fungi, particularly during the stages of grape cultivation. Such influences could determine grape quality (e.g. spoilage, mycotoxin presence), and the diversity of species that enter the grape juice/must for fermentation. Thus, winemaking is a process that involves a diversity of microbial interactions along the production chain. Bacteria make significant contributions to these interactions. Consequently, it is important to have more detailed information about their occurrence and development on grapes.

G G 54

Although the bacterial species associated with wine production have been reported over many years (Amerine and Kunkee 1968; Fleet 1993; Fugelsang 1997), most investigations have focused on target bacterial species such as lactic acid bacteria

(Kunkee 1967, 1974; Wibowo et al. 1985; Lonvaud-Funel 1999) and acetic acid bacteria (Drysdale and Fleet 1988), particularly during the wine fermentation process.

There have been very few analyses of grapes in the vineyard or analyses of bacteria other than lactic and acetic acid bacteria. Moreover, there has been little systematic study of the viticultural and other environmental factors that might affect the bacterial species associated with grapes. It is not unusual to find damaged berries among healthy, undamaged berries on bunches of wine grapes, and it is now acknowledged that the two types of berries will have different ecology that needs to be considered in the overall microbiological assessment.

Current understanding of the microbial ecology of grapes is based upon the use of standard cultural procedures for bacterial isolation and identification. Culture- independent, molecular methods based on the analysis of extracted DNA are now available and offer an alternative approach to profiling the microbial ecology of natural habitats. One of these methodologies is PCR-denaturing gradient gel electrophoresis

(PCR-DGGE), and it is finding increased application to food, beverage and natural ecosystems (Heuer and Smalla 1999; Muyzer 1999; Smit et al. 2001; van Beek and

Priest 2002; Ercolini 2004; Giraffa 2004). In addition to speed and convenience of analysis, PCR-DGGE increases the prospect of detecting viable, but non culturable organisms and giving a more reliable and comprehensive indication of the ecology

(Kuske et al. 1997; Garbeva et al. 2001; Dilly et al. 2004).

G G 55

In this Chapter, the populations and identification of bacterial species associated with wine grapes throughout their cultivation at several Australian vineyards are reported.

Influences of berry damage, vineyard region, pesticide application and seasonal properties on the bacterial ecology are also reported. Bacterial species associated with the grapes were determined by cultural isolation and by PCR-DGGE analysis of extracted DNA.

3.2 MATERIALS AND METHODS

3.2.1 Grape samples

Grapes, collected during October 2001-February 2002 and October 2002-February 2003, were obtained from six commercial vineyards located in the upper and lower Hunter

Valley, Mudgee and Griffith regions of New South Wales, Australia. The northern region (upper and lower Hunter Valley), the northwestern region (Mudgee), and the southwestern region (Griffith) are geographically distinct from each other, representing a spectrum of climates across the regions (i.e. warm and humid Hunter Valley, cooler climate of Mudgee, and the hotter, drier climate of Griffith) (www.southcorp.com.au

2005). These regions are separated from Sydney by distances of approximately 165-595 km (Fig. 3.1). Samples including three varieties of red grapes (Cabernet Sauvignon,

Merlot, Shiraz) and three varieties of white grapes (Chardonnay, Sauvignon Blanc,

Semillon) were collected for examination (Table 3.1). Undamaged, healthy grapes were taken at five phenological stages of development throughout cultivation as shown in

Table 3.2 and Figs. 3.2 and 3.3. Each sample consisted of small clusters or bunches of healthy, undamaged grapes (200 g) that were aseptically removed from at least five different vines within the vineyard and combined to give a total of about 1 kg. Duplicate samples were taken for each grape variety.

G 56

New South Wales Upper Hunter Valley (276km) Lower Hunter Valley Mudgee a (260 km) (165 km)

Griffith Sydney (595 km)

Kilometres (X100) 0123

a, distance from Sydney

Fig. 3.1 Location of vineyard regions in New South Wales (NSW), Australia

Table 3.1 Sources of wine grapes used for the analysis of bacteria

Code Vineyard Vineyard location Grape variety

A Rosemount Upper Hunter Valley, NSW Shiraz, Cabernet Sauvignon, Estate Chardonnay, Semillon

B Fernandez Upper Hunter Valley, NSW Shiraz, Chardonnay

C Lindemans Lower Hunter Valley, NSW Shiraz, Semillon

D Combandry Mudgee, NSW Shiraz, Chardonnay

E Hill of Gold Mudgee, NSW Cabernet Sauvignon, Merlot, Sauvignon Blanc

F McWilliams Griffith, NSW Shiraz, Chardonnay 57

Table 3.2 Stages of cultivation and maturity when grapes were sampled for bacterial analysis

Weeks Maturity stage Description of grapesa before harvest code 14-16 0 Inflorescence and flower formation before bloom (before flower caps loosening), clusters are closed 10-12 I Berries hard and green; about 2-5 mm in diameter, accompanied by closure of bunches 8-10 II Berries, 5-10 mm in diameter (pea size), (Veraison) change colour, berries begin to soften 4-8 III Berries, 8-10 mm in diameter, commencement of ripening

2-4 IIIa Berries ripening 1-2 IV Berries, 10-15 mm in diameter 0 V Berries fully ripe, 10-15 mm in diameter and progress to senescence after this stage a Pratt 1971; Coombe 1992, 1995; Fleet et al. 2002

For each grape variety, samples of damaged grape berries were also collected and examined. Damaged grape berries were collected for two phenological stages only.

These were at the time of commercial harvest and at 2-3 weeks before harvest.

Damaged grapes were identified on the basis of their physical appearance (broken skin, shrivelled, discoloured).

Samples were stored at 4 C and transported to laboratories at UNSW by road or air.

Grapes were analysed within 24 h of harvest from the vine. 58 I II

10-12 weeks before harvest 8-10 weeks before harvest (veraison)

III IV

4-8 weeks before harvest 1- 2 weeks before harvest

V V

harvest (damaged grapes) harvest

Fig. 3.2 Maturity stages of Shiraz grapes examined for bacteria (also see Table 3.2). 59 I II

10-12 weeks before harvest 8-10 weeks before harvest (veraison)

III IV

4-8 weeks before harvest 1- 2 weeks before harvest

V V

harvest (damaged grapes) harvest

Fig. 3.3 Maturity stages of Chardonnay grapes examined for bacteria (also see Table 3.2) G 60

3.2.2 Analysis of bacteria on grapes

Bacteria on grapes were determined by (i) direct culture by plating on agar media, (ii) enrichment culture in liquid media followed by plating on agar media, and (iii) PCR-

DGGE analysis, as outlined in Fig. 3.4. Individual grape berries were randomly and aseptically removed from each cluster or bunch to give a composite sample of about 20 g (maturity stage I, Figs. 3.2, 3.3) and 50 g (maturity stages II – V, Figs. 3.2, 3.3). They were suspended in 180 mL or 450 mL, respectively, of sterile 0.1% Bacteriological

Peptone (Oxoid, Melbourne, Australia) solution, containing 0.01% Tween 80 (Sigma, St.

Louis, MO.) in a conical flask, and orbitally shaken at 180 rpm for 30 min at 25qC. The rinse was poured from the grapes and examined for bacteria by both cultural and PCR-

DGGE methods.

3.2.3 Culture on agar media

Rinses from the grape samples were serially diluted in 0.1% Bacteriological Peptone solution and duplicate samples (0.1 mL) of each dilution were spread inoculated onto the surface of plates of Plate Count Agar (PCA, Oxoid), de Man Rogosa and Sharpe agar (MRS agar, Oxoid) and MRST agar. MRST agar consisted of MRS agar supplemented with 15% tomato juice (Berri, Australia) and adjusted to pH 5.5 with HCl.

Plate count agar was used to isolate the total and general bacterial species occurring on the surface of wine grapes, and MRS and MRST agars were used for the isolation of lactic acid bacteria. Fortification of MRS with tomato juice has been reported to improve the isolation of lactic acid bacteria associated with wines (Pan et al. 1982). All media were supplemented with cycloheximide (Sigma) to a final concentration of 100 mg/L to prevent the growth of yeasts and other fungi. Plates were incubated at 25qC for

2-7 days until prominent colonies were observed. Representatives of the different types

G 61

Grape berries

Enrichment culture analysis Diluent 450 mL

Shaking 50 g 30 min 10 g berries 10 g berries + + Nutrient broth MRS broth (0.1% Bacteriological peptone (90 mL) (90 mL) + Tween 80)

Incubation 25 C / 1~5 days Supernatant

Agar culture analysis Molecular analysis Diluting samples

Sediment microbial cells

DNA Extraction

Plating PCR amplification of 16S rDNA Plate Count Agar (PCA), MRS agar, MRST agar DGGE

Incubation (25 C / 2~7 days) Sequencing bands Colony counts and identification of colonies Species identity by DNA sequencing

Fig. 3.4 Outline of protocol for the analysis of bacteria on wine grapes G 62 of colonies were selected and restreaked at least three times to obtain pure cultures.

They were identified by observation of cellular morphology, Gram staining, catalase and oxidase tests and by 16S ribosomal DNA sequence analysis.

3.2.4 Enrichment culture

Grapes (10 g) were aseptically transferred to flasks containing 90 mL of Nutrient broth

(Oxoid) and MRS broth (Oxoid), respectively and incubated at 25qC for 1-5 days or until visibly turbid. All media were supplemented with 100 mg/L of cycloheximide

(Sigma) to prevent the growth of yeasts and fungi. Cultures were sampled, diluted as required in 0.1 % Bacteriological Peptone solution, and aliquots (0.1 mL) were spread inoculated over the surfaces of the corresponding agar media. After incubation at 25qC for 1-5 days, dominant and representative colonies from Nutrient agar and presumptive lactic acid bacteria (Gram-positive rods or cocci, catalase negative) from MRS agar were noted and isolated for identification.

3.2.5 DNA extraction for bacterial identification and PCR-DGGE analysis

Microbial pellets from cell cultures (2 mL pure culture sample, centrifuged at 12,000ug,

10 min, 4qC) and from the rinses of grapes (approx. 180 mL or 450 mL, centrifuged at

16,000ug, 15 min, 4qC) were prepared, and the DNA was extracted as described by

Lopez et al. (2003). The microbial pellet from cell cultures was suspended in 200 Pl of breaking buffer (2% Triton X-100, 1% SDS, 100 mM NaCl, 10 mM Tris-Cl [pH 8.0], 1 mM EDTA [pH 8.0]) containing 0.3 g of zirconia/silica beads (0.1 mm diameter,

BioSpec Products Inc., Bartlesville, OK). The cells were homogenized twice for 40 s at

4,600 rpm in a bead beater (Mini-BeadBeater-1•, BioSpec Product Inc.) in the

G G 63 presence of 200 PL of phenol/choloroform/iso-amylalcohol (25:24:1). Two hundred microliters of TE buffer (10 mM Tris-Cl, 1 mM EDTA, pH 8.0) were then added, and the bead cell mixture was centrifuged at 12,000ug for 10 min at 4qC. The aqueous phase was transferred to another microcentrifuge tube, and the DNA was precipitated by the addition of 0.6 volume of isopropanol and recovered by centrifugation at 12,000ug for

10 min at 4qC. The DNA pellet was washed with 70% ethanol, dried and resuspended in

50 PL of sterile TE buffer for identification by 16S ribosomal DNA sequencing analysis.

DNA in the sedimented microbial pellets from rinses of grapes was extracted by the same procedure as described above with the following modification. The supernatant of the phenol/chloroform/isoamylalcohol extracts was transferred to DNeasy kit (Qiagen

Pty. Ltd., Clifton Hill, Australia) and purified according to the protocol supplied by the manufacturer. In the final purification step, DNA was totally eluted from the filter with

150 PL of RE buffer supplied in the kit. This DNA was used for PCR-DGGE analysis.

3.2.6 PCR amplification of DNA for sequence identification and PCR-

DGGE analysis

Purified DNA from pure cultures was amplified with universal primers flanking the region between 968 and 1401 (E. coli numbering) for identification of the bacteria by

16S ribosomal DNA sequence analysis as described by Bae et al. (2004). All primers used in this study were obtained from SigmaGenosys, Australia. The PCR mix contained 1uPCR buffer (10mM Tris-HCl [pH 8.3], 50 mM KCl), 0.2 PM of each primer, 200 PM of each dNTP (Roche Diagnostics, Indianapolis, IN, USA), 1.5 mM

MgCl2, 1.25 U of Gold Taq DNA polymerase (AmpliTaq•, Roche Molecular Systems,

G G 64

Brachburg, NJ, USA) and 10ng of purified template DNA in 50 PL final volume.

Amplification was carried out with a 9600 Thermal Cycler (Applied Biosystems) with the following program: initial denaturation at 94qC for 7 min, 10 cycles at 94qC for 30 s,

50qC for 30 s, and 72qC for 45 s, followed by 20 times the same cycle with each successive cycle at 5 s longer elongation time. The final elongation was conducted for

10 min at 72qC. This PCR amplicon was used for sequence analysis.

To investigate the bacterial communities by PCR-DGGE, two sets of primers were employed. Purified DNA extracted from rinses of grapes was amplified with primers

341f-gc/518r (Muyzer et al. 1993) and 968f-gc/1401r (Nübel et al. 1996), respectively.

The 40 nucleotide GC rich sequence (5’-cgc ccg ccg cgc ccc gcg ccc gtc ccg ccg ccc cgc ccg g-3’) is attached to the 5’ end of forward primers (341f and 968f) to facilitate the detection of sequence variations of amplified DNA fragments by subsequent DGGE

(Myers et al. 1985; Muyzer et al. 1993). The first primer set, 341f-gc (5’- gc clamp-

CCT ACG GGA GGC AGC AG-3’) and 518r, (5’-ATT ACC GCG GCT GCT GG-3’), targets the V3 region of the 16S ribosomal RNA (Muyzer et al. 1993). Reaction mixtures were as described above with the addition of 0.25 Pg of T4 gene 32 protein

(Roche Diagnostics GmbH, Germany). T4 gene 32 protein, a single-strand binding protein, was added to increase amplification efficiency (Schwieger and Tebbe 1997).

Amplification was performed under the following program: initial denaturation at 95qC for 7 min, 20 cycles at 94qC for 1 min (denaturation), 65qC for 1 min (annealing) and

72qC for 1min (extension) with a 1qC touchdown every second cycle, followed by 10 cycles with an annealing temperature of 55qC, and the final extension was conducted for 10 min at 72qC. A second pair of primers spanning the V6-V9 region of the 16S

G G 65 rRNA gene, 968f-gc (5’-gc clamp- AAC GCG AAG AAC CTT AC-3’) and 1401r (5’-

CGG TGT GTA CAA GAC CC-3’) were also used (Nübel et al. 1996). The PCR mixture was modified from that of the first primer pair by increasing the MgCl2 concentration to 3.75 mM. Amplification was performed with the reaction mixtures as described above under the following program: initial denaturation at 95qC for 7 min, 10 cycles at 94qC for 1 min (denaturation), 60qC for 1 min (annealing) and 72qC for 2 min

(extension) with a 1qC touchdown every second cycle, followed by 20 cycles with an annealing temperature of 55qC, and the final extension was conducted for 10 min at

72qC. The PCR amplicons were confirmed by 2% agarose gel electrophoresis.

3.2.7 Denaturing gradient gel electrophoresis (DGGE)

The resultant PCR amplicons from Section 3.2.6 were resolved by DGGE using a

DCode Universal Mutation Detection System (Bio-Rad, Hercules, CA, USA).

Essentially, DGGE was performed as described by Muyzer et al. (1993). Eight percent polyacrylamide gels (acrylamide:N,N’-methylene bisacrylamide ratio, 19:1, Bio-Rad) with 30-60% linear denaturing gradient (100% denaturant corresponds to 7 M urea and

40% formamide) were used for analysing amplicons of 341f-gc/518r. Electrophoresis was performed for 30 min at 20 V and for 6 h at 120 V with a constant temperature at

60qC in 1u TAE buffer (40 mM Tris-acetate, 1mM EDTA). Six percent polyacrylamide gels with 45 - 65% linear denaturing gradient were used for analysing amplicons produced with primers of 968f-gc/1401r, and electrophoresis was run for 30 min at 20 V, then for 14 h at 100V. Subsequently, gels were stained with SYBR“ Green I

(Amresco“, Ohio, USA) for 20 min in 1u TAE buffer (40 mM Tris-acetate, 1 mM

EDTA, pH 8.0) and photographed under UV transillumination with a Polaroid DS-34

G G 66 camera.

DNA bands in DGGE gels were excised and incubated overnight in 50 ȝL of sterile

Milli-Q water at 4qC. Eluted DNA was reamplified with corresponding primers and the

PCR products were re-evaluated by DGGE. Only products that migrated as a single band and at the same distance with respect to the original sample were then excised and amplified with corresponding primers without a GC clamp. Amplified products were purified with QIAquick PCR purification kit (Qiagen Pty Ltd., Clifton Hill, Australia), and subjected to DNA sequence analysis.

3.2.8 Construction of DGGE validation marker

To verify the correctness of the applied gradient and to allow comparison of the DGGE banding patterns between samples on gels, DGGE validation markers were constructed with bacterial isolates from wine grapes. The markers (M) were generated by pooling the amplicons produced with either 341f-gc/518r primers or 968f-gc/1401r primers.

Good separation of DNA bands was obtained after PCR amplified with 341f-gc/518r primers as shown in Fig. 3.5. Amplicons of some bacterial species produced by primers of 968f-gc/1401r were resolved as multiple bands on DGGE gels regardless of different conditions used in preparation or running of the electrophoresis gel. Therefore, only

341f-gc/518r primers were chosen for analysis of bacterial ecology by DGGE. While all the isolates gave one prominent DNA band, it should be noted that three weaker bands were also obtained for the DGGE profile of B. thuringiensis.

G 67

MM12345678

Fig. 3.5 DGGE of the 16S rDNA amplicons of bacterial isolates from wine grapes. Lanes: M, mixture of species in lanes 1-8; 1, Arthrobacter gandensis (AJ316140); 2, Rhodococcus equi (AF490539); 3, Curtobacterium flaccumfaciens (AY273208); 4, Bacillus thuringiensis (AF155955); 5, Microbacterium paraoxydans (AJ491806); 6, Lactococcus lactis (AF493058); 7, Pantoea agglomerans (AJ233423); 8, Lactobacillus plantarum (AL935261).

3.2.9 DNA sequencing

PCR amplicons of DNA from bacterial isolates and from band eluates of DGGE gels were identified by sequence analysis of the 16S ribosomal DNA and used directly for sequencing with the ABI PRISMR BigDyeTM Terminator v3.1 Cycle Sequencing kit

Applied Biosystems, Foster City, CA, USA). The products were analysed for their sequence at the Automated DNA analysis facility, School of Biotechnology and

Biomolecular Sciences, University of New South Wales. The sequences were aligned to determine the closest known relatives of partial 16S rDNA gene fragments available from BLAST in the GENBANK database (Karlin and Altschul 1990). G 68

3.2.10 Species identification

All isolates were examined for cell morphology and Gram stain, catalase and oxidase reactions according to standard procedures (Smitbert and Krieg 1994) and correlated with sequence data. Isolates were identified by sequence data. Homology data and accession numbers for the different species and samples analysed are shown in

Appendix 1. Also, DNA base sequences from DNA bands were aligned to determine the closest known relatives of partal 16S rDNA and shown in Appendix 2.

3.3 RESULTS

3.3.1 Climatological data for the vineyard regions in the years of 2001-

2003

Fig. 3.6 shows the rainfall and temperature data of the vineyard regions in the period of the grape season for 2001-2002 and 2002-2003, as well as those for the overall period

2000-2004, which was used for reference and standardisation purposes. These data were supplied by the Australian Government Bureau of Meteorology. Arrows indicate the time when wine grapes were sampled. The rainfall of the upper Hunter Valley region in the grape growing period of 2001-2002 was consistent with yearly average throughout cultivation, but was unusually heavy three days before the sampling date at harvest time.

For the season of 2002-2003, three of the vineyards (Upper Hunter, Lower Hunter and

Mudgee region) generally experienced much lower rainfall than average. This was not the case for the Griffith region (Fig. 3.6d) which represents a consistently drier region, with higher average temperature. In most cases, temperatures during the 2002-2003 season were higher than average, especially during the October-November period, just before starting of veraison. On some days during November and December,

G 69

temperatures in the vineyards reached 45 C.

Rainfall Temperature 180 40 (a) (a) 150 30 120

90 20

60

Temperature (10 C) 30

180 40 (b) (b) 150 30 120

90 20

Rainfall (mm)60 Rainfall (mm) 10 30 Temperature ( C)

180 40 (c) (c) 150 30 120

90 20

Rainfall (mm) 60 10 Temperature ( C) 30

180 40 (d) (d) 150 30 120

90 20

Rainfall (mm) 60 10 Temperature ( C) 30

0 0 Aug Sep Oct Nov Dec Jan Feb Aug Sep Oct Nov Dec Jan Feb Month Month

Fig. 3.6 Rainfall (mm) and mean daily maximum temperature ( C) of vineyard regions during the period of grape growth (a) Upper Hunter Valley region, 2001-2002 and 2002-2003; (b) Lower Hunter Valley region, 2002-2003; (c) Mudgee region, 2002-2003; (d) Griffith region, 2002-2003. Arrows indicate time when wine grapes were sampled ( , ). ( ), Mean rainfall (mm) during 2000-2004; ( ), Mean daily maximum temperature during 2000-2004; (,,),2001-2002; (,,),2002-2003. G 70

3.3.2 Populations of bacteria on wine grapes during cultivation

To examine the population and diversity of bacterial species occurring on wine grapes, six different grape varieties were sampled at five phenological stages throughout cultivation from six commercial vineyards for the 2002-2003 season. Vineyard A

(Upper Hunter Valley) was also examined throughout the 2001-2002 season. No lactic acid bacteria were detected on MRS or MRST agar plates for grapes of vineyard A during 2001-2002. Instead, colonies of Bacillus thuringiensis were prevalent on these media, as well as on PCA. The total bacterial populations for the grapes were taken from PCA. A mixture of bacteria, including B. thuringiensis, generally grew on plates of

MRS and MRST agar, but the populations were 10-100 fold less than those on PCA.

Lactic acid bacteria (Gram positive, catalase negative) were detected on these plates on only three occasions during 2002-2003 at populations around 102-103 CFU/g (Cabernet

Sauvignon, maturity stage II, vineyard A; Shiraz and Semillon, maturity stage I, vineyard C).

Fig. 3.7 shows the total populations of bacteria on several grape varieties (healthy, undamaged grape berries only) during cultivation. The populations were the mean counts of bacteria detected from duplicates of grape samples plated in duplicate on PCA.

The populations varied between 102-106 CFU/g, with higher populations being found at the early stages of cultivation. At the time of harvest, bacterial populations on grapes were generally about 102-103 CFU/g except for grapes from vineyard A in 2001-2002, where the populations were approximately 103-104 CFU/g. There were no consistent influences of grape variety on bacterial populations. Grapes from vineyard A, tended to have slightly higher populations, especially during the wetter season of 2001-2002.

G 71

7 7 Vineyard A, 2001-2002 (a) Vineyard A, 2002-2003 (b) cs 6 s 6 sm c 5 sm 5

4 4 cs s c 3 3 Log CFU/g

2 2

1 1

0 0 IVIIIII IV III III IV V

7 7 Vineyard B, 2002-2003 (c) Vineyard C, 2002-2003 (d) 6 6 s sm

5 5

4 4

3 3

Log CFU/g s c 2 2

1 1

0 0 II III IV V IVII III IV

7 7 m Vineyard D, E, 2002-2003 (e) Vineyard F, 2002-2003 (f) 6 6 c

5 sb 5 s s

c 4 4 cs

3 3 Log CFU/g 2 2

1 1

0 0 IVIIIII IV III III IV V Phenological stage of berry development Phenological stage of berry development

Fig. 3.7 Total populations of bacteria on grapes during cultivation at vineyards in different regions of New South Wales (a) vineyard A, 2001-2002, (b) vineyard A, 2002-2003, (c) vineyard B, 2002-2003, (d) vineyard C, 2002-2003, (e) vineyards D and E, 2002-2003, (f) vineyard F, 2002-2003. Phenological stages of berry development are given in Table 3.2. Shiraz (s, ); Cabernet Sauvignon (cs, ); Merlot (m, ); Chardonnay (c, ); Semillon (sm, ); Sauvignon Blanc (sb, ). 72

Fig. 3.8 shows the mean counts of bacteria on grapes, according to red and white grape

varieties from different regions. For red wine grapes, the bacterial populations were

more variable at the earlier than later stages of cultivation. The populations on the

grapes decreased and showed less variation as the berries matured to harvest. Similar

trends were observed for the white grape varieties. Both types of grapes exhibited

bacterial populations of 1023 -10 CFU/g at harvest time. Even though there was higher

rainfall during the 2001-2002 season than during the 2002-2003 season, and the

vineyard F was drier than the other regions, the total bacterial populations on healthy

red or white undamaged berries at the time of harvest were about 1023 -10 CFU/g.

7 7 (a) Red grapes (b) Red grapes 6 6

5 5

4 4

3 3

2 2 Population (Log CFU/g) 1 1 I II III IV V I II III IV V 7 7 (c) White grapes (d) White grapes 6 6

5 5

4 4

3 3

2 2 Population (Log CFU/g) 1 1 I II III IV V I II III IV V Phenological stage of berry development Phenological stage of berry development Fig. 3.8 Mean counts of bacteria from red and white wine grapes sampled from different vineyards according to maturity (a), (b) mean counts of bacteria of red wine grapes; (c), (d) mean counts of bacteria of white wine grapes. ( ), vineyard A, 2001-2002; ( ), vineyard A, 2002-2003; ( ), vineyard B, 2002-2003; ( ), vineyard C, 2002-2003; ( ), vineyard D, 2002-2003; ( ), vineyard E, 2002-2003; ( ), vineyard F, 2002.

A comparison of bacterial populations for undamaged and damaged grape berries is

shown in Fig. 3.9. Damaged grapes were collected at two times, 1-2 weeks prior to 73

commercial harvest time (maturity stage IV) and at commercial harvest time (maturity

stage V). Generally, damaged grapes gave total bacterial populations about 10-100 fold

more than undamaged grapes. These differences were particularly evident for grapes

harvested from vineyard A during the 2001-2002 season, when there were significant

rainfalls around the time of harvesting.

(a) maturity stage IV, undamaged (b) maturity stage IV, damaged

c c F s s

sb sb E m m cs cs * c c D s * 2002- s 2003 sm sm* C s s* c c B s s sm sm c c A cs cs s s * sm sm c *c 2001-2002, A cs cs * s s *

(c) maturity stage V, undamaged (d) maturity stage V, damaged

c c F s s *

sb sb E m m * cs cs 2002- c c D s s 2003 sm sm C s s

c c B s s * sm sm c c A cs cs s s sm sm c c * 2001-2002, A cs cs * s s* * 0 1 2 3 4 5 6 061 2 3 4 5 Log CFU/g Log CFU/g Fig. 3.9 Populations of bacteria on undamaged and damaged grapes at maturity stages IV and V from different vineyards A-F, during the 2001-2002 and 2002-2003 seasons (a) undamaged grapes at maturity stage IV; (b) damaged grapes at maturity stage IV; (c) undamaged grapes at maturity stage V; (d) damaged grapes at maturity stage V. Shiraz (s, ); Cabernet Sauvignon (cs, ); Merlot (m, ); Chardonnay (c, ); Semillon(sm, ); Sauvignon Blanc (sb, ). A t-test was conducted and asterisks (* ) indicate significant differences between undamaged and damaged samples at a 95% level. G 74

3.3.3 The diversity of bacterial species associated with wine grapes during cultivation

Six different grape varieties from several vineyards in NSW (Table 3.1) were routinely examined for bacteria throughout cultivation from October 2001 to February 2002 and from October 2002 to February 2003. For the 2001-2002 season, grape samples were simultaneously examined by plate culture and PCR-DGGE methods. With grapes from the 2002-2003 season, enrichment culture was also performed as an additional assay.

3.3.3.1 Hunter Valley region, 2001-2002

Bacillus thuringiensis was the most prevalent species on every grape variety collected from the vineyard A, during the 2001-2002 season (Table 3.3). Its population was highest (104-106 CFU/g) at the early stages of grape development and decreased to 102-

104 CFU/g as the grape matured at harvest. The prevalence of B. thuringiensis on the grapes was also demonstrated by PCR-DGGE analysis except on Semillon grapes

(maturity IV) (Fig. 3.10).

After B. thuringiensis, Curtobacterium flaccumfaciens, and species of Microbacterium,

Pantoea and Pseudomonas were frequently isolated. Curtobacterium flaccumfaciens was consistently found on mature grapes (stage V) of each variety by both plate culture and DGGE methods, but was inconsistently seen at earlier stages. On some occasions, it was the most prevalent bacterial species on grapes at the time of harvest. Other bacterial species were inconsistently isolated from grapes throughout cultivation. Some of these species were only detected by PCR-DGGE and not by plate culture (e.g. Brevibacterium,

Pseudomonas, Ralstonia, Rhodocista, Sphingomonas).

G G 75

Table 3.3 Bacterial species and populations (CFU/g) on grapes at different stages of maturity in vineyard A during the 2001-2002 season as determined by (a) plate culture and (b) PCR-DGGE

Grape Detection Maturity of grapes variety method I II III IV V Shiraz a B. thuringiensis (5.6u105) B. thuringiensis (6.4u104) B. thuringiensis (5.1u104) B. thuringiensis (4.1u102) B. thuringiensis (5.5u102) C. flaccumfaciens (6.8u102) b B. thuringiensis B. thuringiensis B. thuringiensis B. thuringiensis B. thuringiensis Ps. alcaligenes Ag. tumefaciens Br. equis Ralstonia sp. C. flaccumfaciens Rhodocista sp. Mb. imperiale Cabernet a B. thuringiensis (7.5u105) B. thuringiensis (1.0u105) B. thuringiensis (2.5u103) B. thuringiensis (1.8u103) B. thuringiensis (1.3u103) Sauvignon Microbacterium sp. (2.3u102) C. flaccumfaciens (3.0u102) C. flaccumfaciens (2.5u103) b B. thuringiensis B. thuringiensis B. thuringiensis B. thuringiensis B. thuringiensis Pseudomonas sp. Sphingomonas sp. C. flaccumfaciens Bacterium PSD-1-6 Chardonnay a B. thuringiensis (9.9u104) B. thuringiensis (2.8u104) B. thuringiensis (2.9u104) B. thuringiensis (1.2u103) B. thuringiensis (1.3u103) C. flaccumfaciens (4.0u104) C. flaccumfaciens (2.4u103) C. flaccumfaciens (3.4u103) Pseudomonas sp. (1.4u103) b B. thuringiensis B. thuringiensis B. thuringiensis B. thuringiensis B. thuringiensis Pseudomonas sp. C. flaccumfaciens C. flaccumfaciens Sphingomonas sp. Sphingomonas sp. Semillon a B. thuringiensis (3.9u104) B. thuringiensis (2.7u104) B. thuringiensis (3.0u103) B. thuringiensis (3.2u102) B. thuringiensis (2.0u102) Microbacterium sp. (1.1u102) Microbacterium sp. (3.0u102) C. flaccumfaciens (1.0u103) Pantoea sp. (2.0u103) b B. thuringiensis B. thuringiensis B. thuringiensis Ar. gandensis B. thuringiensis C. flaccumfaciens Pantoea sp. Ps. putida Sphingomonas sp.

G Shiraz Cabernet Sauvignon Chardonnay Semillon M I II III IV VM I II III IV V MM I II III IV V I II III IV V Bac

Ps a

Ps Cf Cf Cf Bt Bt Ps Cf Bt Cf Bt Sphin Ps p Vv Cf Ag t Sphin Pa Br e Vv Cf Sphin Vv Cf Rals Mim Pa

Ar g Rhod

Fig. 3.10 PCR-DGGE analysis of bacterial DNA in rinses of wine grapes from vineyard A at different stages during cultivation in 2001-2002. Lanes: I, II, III, IV, V, represent the stages of maturity referred to in Table 3.2. M, marker referred to in Fig. 3.5. Ag t, Ag. tumefaciens; Ar g, Ar. gandensis; Br e, Br. equis; Bt, B. thuringiensis; Cf, C. flaccumfaciens; Mim, Mb. imperiale; Pa, Pantoea sp.; Ps, Pseudomonas sp.; Ps a, Ps. alcaligenes; Ps p, Ps. putida; Rals, Ralstonia sp.; Rhod, Rhodocista sp.; Sphin, Sphingomonas sp.; Bac, Bacterium PSD-1-6; Vv, Vitis vinifera. 76 G 77

3.3.3.2 Hunter Valley region, 2002-2003

Vineyard A

For grapes collected at vineyard A in 2002-2003, B. thuringiensis was the most prevalent species found on all varieties, and throughout the different stages of berry development (Table 3.4). Its prevalence was confirmed by all three analytical methods- plate culture, enrichment culture and PCR-DGGE (Table 3.4, Fig. 3.11). Its population was relatively low, at approximately 102-103 CFU/g, except for Semillon grapes where it occurred at 104-105 CFU/g during the early stages of berry development. Unlike the

2001-2002 data for these grapes, C. flaccumfaciens was not frequently isolated, but was found on Chardonnay grapes at harvest.

Various other bacterial species within the genera Acidovorax, Arthrobacter, Bacillus,

Brachybacterium, Curtobacterium, Erwinia, Flavobacterium, Hafnia, Pantoea,

Pseudomonas, Serratia and Sphingomonas were detected infrequently by plate culture, especially at maturity stage I. Generally, their populations were low, at 102-103 CFU/g.

Only one sample (Chardonnay) showed the presence of some of these species at the stage of commercial harvest. Most of these species were not detected by PCR-DGGE

(Table 3.4). Although PCR-DGGE failed to detect some bacterial species that were found by plate culture, it did reveal the presence of several species that were not found by plate culture. These were species of Deinococcus, Flavobacterium, Paenibacillus,

Pseudomonas, Ralstonia, Sphingomonas, Staphylococcus and Streptosporangium. The repeated detection of Sphingomonas sp. in Cabernet Sauvignon grapes is notable (Fig.

3.11). Lactic acid bacteria were rarely isolated, even by enrichment culture. Lactococcus lactis (Shiraz, maturity stage I) and Enterococcus faecalis (Cabernet Sauvignon, maturity stage II) were the only lactic acid bacteria found, and they were isolated by

G G 78

Table 3.4 Bacterial species and populations (CFU/g) on grapes at different stages of maturity in vineyard A during the 2002-2003 season as determined by (a) plate culture, (b) PCR-DGGE and (c) enrichment culture

Grape Detection Maturity of grapes variety method I II III IV V a B. thuringiensis (1.0u103) B. thuringiensis (1.0u102) B. thuringiensis (5.0u102) B. thuringiensis (2.0u102) B. thuringiensis (2.0u102) B. megaterium (3.5u102) C. flaccumfaciens (4.0u102) Ar. luteolus (5.0u102) Pa. ananatis (1.0u102) Acidovorax sp. (3.5u102) S. liquefaciens (2.5u102) Shiraz b B. thuringiensis B. thuringiensis B. thuringiensis B. thuringiensis B. thuringiensis Pseudomonas sp. Ps. viridiflava c Lc. lactis B. thuringiensis B. thuringiensis B. thuringiensis B. thuringiensis Br. acetylicum B. pumilus Yersinia sp. B. thuringiensis a B. thuringiensis (1.0u102) B.thuringiensis (1.0u102) B. thuringiensis (9.5u102) B. thuringiensis (3.5u102) B. thuringiensis (2.0u102) C. flaccumfaciens (3.5u102) E. faecalis (4.5u102) Sphingomonas sp. (5.0u102) Br. paraconglomeratum Pantoea sp. (5.0u102) (4.5u102) Er. cypripedii (3.0u103) Ps. aureofaciens (9.5u102) Cabernet b ND B. thuringiensis B. thuringiensis B. thuringiensis Sphingomonas sp Sauvignon Sphingomonas sp. Sphingomonas sp. Uncultured bacterium clone IG6 (Un) Ralstonia sp. c B. thuringiensis B. thuringiensis B. thuringiensis B. thuringiensis B. thuringiensis Acinetobacter sp. B. sphaericus Br. acetylicum

G G 79

Table 3.4 (Continued) Bacterial species and populations (CFU/g) on grapes at different stages of maturity in vineyard A during the 2002- 2003 season as determined by (a) plate culture, (b) PCR-DGGE and (c) enrichment culture

Grape variety Detection Maturity of grapes method I II III IV V a B. thuringiensis (6.5u102) B.thuringiensis (1.2u104) B. thuringiensis (2.1u103) B. thuringiensis (8.5u102) B. thuringiensis (4.5u102) B. megaterium (1.0u102) Arthrobacter sp. (1.0u102) C. flaccumfaciens (1.0u102) C. flaccumfaciens (1.5u102) Flavobacterium sp. (1.0u102) Pseudomonas sp. (4.5u102) Hafnia alvei (2.5u102) Chardonnay b ND B. thuringiensis B. thuringiensis B. thuringiensis B. thuringiensis Mb. paraoxydans Ps. putida Ps. viridiflava Pseudomonas sp. c B. thuringiensis B. thuringiensis B. thuringiensis B. thuringiensis B. thuringiensis Mb. paraoxydans a B. thuringiensis (1.0u105) B. thuringiensis (1.5u104) B. thuringiensis (8.0u102) B. thuringiensis (1.4u103) B. thuringiensis (5.5u102) Arthrobacter sp. (1.0u102) b B. thuringiensis B. thuringiensis B. thuringiensis B. thuringiensis B. thuringiensis Bacterial species Flavobacterium sp. D. radiodurans 16S rRNA gene Pb. dendritiformis Ps. viridiflava Staphylococcus sp. Ps. viridiflava Semillon Uncultured bacterial isolates Streptosporangium sp. Uncultured earthworm intestine bacterium clone c B. thuringiensis B. thuringiensis B. thuringiensis B. thuringiensis B. thuringiensis Br. acetylicum B. mycoides Mb. phyllosphaerae Sporo. inulinus ND, Not detected

G Shiraz Cabernet Sauvignon Chardonnay Semillon M I II III IV VMM I II III IV V I II III IV V M I II III IV V

Ps Ps v Staph Ps Ps p Ps Ps v Ps v Ps Un

Bt Flav Bt Sphin Bt Bt Un Vv Vv Vv Vv Rals P den Mpa D rad Un Un Stre

Fig. 3.11 PCR-DGGE analysis of bacterial DNA in rinses of wine grapes from vineyard A at different stages during cultivation in 2002-2003. Lanes: I, II, III, IV, V, represent the stages of maturity referred to in Table 3.2. M, marker referred to in Fig. 3.5. Bt, B. thuringiensis; D rad, D. radiodurans; Flavo, Flavobacterium sp.; Mpa, Mb. paraoxydans; P den, Pb. dendritiformis; Ps, Pseudomonas sp.; Ps p, Ps. putida; Ps v, Ps. viridiflava; Rals, Ralstonia sp.; Sphin, Sphingomonas sp.; Staph, Staphylococcus sp.; Stre, Streptosporangium sp.; Un, Uncultured bacterial isolates; Vv, Vitis vinifera. 80 G 81 enrichment culture and plate culture, respectively (Table 3.4). They were not detected by PCR-DGGE analysis.

Vineyard B

Grapes in vineyard B (Upper Hunter Valley, NSW, about one kilometer from vineyard

A), were cultivated without the routine application of either chemical (e.g. Thiovit Jet,

Delan 700 DF, Kocide Blude, Oxydul) or biological pesticides (e.g. BT, Dipel Forte,

Delfin WG). Vineyard B, is a commercial “organic” vineyard, and no pesticides had been applied in its environment for many years. Consequently, it was of interest to examine the bacterial ecology of grapes from this vineyard.

Table 3.5 shows the bacterial species and their populations found on Shiraz and

Chardonnay grapes harvested from vineyard B. Plate culture gave very low (101-102

CFU/g) to non-detectable bacterial populations. Bacillus thuringiensis was isolated from samples of both grape varieties at maturity stage IV, but not at the time of commercial harvest (stage V). Curtobacterium flaccumfaciens was also isolated from these grapes at stages III and IV. Neither of these species were detected by PCR-DGGE at these stages. However, enrichment culture demonstrated the presence of B. thuringiensis on all grape samples throughout cultivation. PCR-DGGE revealed the consistent presence of some unusual species not detected by plate and enrichment culture. These were Bacillus benzoevorans and endosymbiont of the insect Brevipalpus lewisi (Fig. 3.12). Brevipalpus lewisi is commonly known as citrus flat mite or bunch mite, and is directly harmful as a crop pest and a vector of the plant virus, citrus leprosies virus. The insect has a very extensive host range, feeding on citrus, grapes, alfalfa, pistachio, fig and many other crops (Balevski et al. 1970; Bournier 1976;

G G 82

Table 3.5 Bacterial species and populations (CFU/g) on grapes at different stages of maturity in vineyard B during the 2002-2003 season as determined by (a) plate culture, (b) PCR-DGGE and (c) enrichment culture G Grape Detection Maturity variety method II III IV V Shiraz a Bacillus sp. (1.5u102) C. flaccumfaciens (2.0u102) C. flaccumfaciens (5.0u101) Pseudomonas sp. (5.0u101) Rhodococcus sp. (1.0u102) B. thuringiensis (5.0u101)

b B. benzoevorans Endosymbiont of B. benzoevorans B. benzoevorans Brevipalpus lewisi Endosymbiont of Lc. lactis Brevipalpus lewisi Endosymbiont of Brevipalpus lewisi c B. thuringiensis B. thuringiensis B. thuringiensis B. thuringiensis Chardonnay a Ar. chlorophenolicus C. flaccumfaciens (5.0u101) B. thuringiensis (5.0u101) ND (1.0u102) Pa. agglomerans. (5.0u101) Ar. gandensis (5.0u101) b B. benzoevorans B. benzoevorans B. benzoevorans B. benzoevorans B. thuringiensis Endosymbiont of Brevipalpus Endosymbiont of Lc. lactis lewisi Brevipalpus lewisi Endosymbiont of Uncultured Methylobacterium Brevipalpus lewisi sp. clone Uncultured bacterium c B. thuringiensis B. thuringiensis E. mundtii B. thuringiensis B. thuringiensis ND, Not detectedG

G 83 http://cals.arizona.edu/crops/citrus/insects/citrusinsect.html). It is common in United

States, Europe, Asia, Africa (Egypt), Oceania, and is categorized as a risk group 1 pest for fresh fruit/vegetables, including the grapes, Vitis vinifera (Buchanan and Amos

1992; Magarey et al. 1999).

Lactic acid bacteria, E. mundtii was found on Chardonnay near the time of commercial harvest (maturity stage IV) by enrichment culture and Lc. lactis were found on Shiraz and Chardonnay samples at the time of commercial harvest by PCR-DGGE, respectively

(Table 3.5, Fig. 3.12).

Shiraz Chardonnay MII IIIIV V MII IIIIV V

Be Be Lc l Lc l Endosymbiont of Brevipalpus lewisi Endosymbiont Bt of Brevipalpus lewisi Vv Vv Un m Un

Fig. 3.12 PCR-DGGE analysis of bacterial DNA in rinses of wine grapes from vineyard B at different stages during cultivation in 2002-2003. Lanes: II, III, IV, V, represent the stages of maturity referred to in Table 3.2. M, marker referred to in Fig. 3.5. Be, B. benzoevorans; Cf, C. flaccumfaciens; Lcl, Lc. lactis; Un m, Uncultured Methylobacterium sp. clone; Un, Uncultured bacterium; Vv, Vitis vinifera. G 84

Vineyard C

Bacillus thuringiensis was the most prevalent species on grapes collected from vineyard

C throughout cultivation, where its population ranged between 101-105 CFU/g (Table

3.6). It was consistently isolated at all stages of berry development by plate and enrichment culture. Its presence was also demonstrated by PCR-DGGE analysis, except for Shiraz and Semillon grapes at maturity stages IV and V, respectively, where the populations were 101-102 CFU/g (Fig. 3.13). Except for the detection of B. thuringiensis,

PCR-DGGE data were not in good agreement with those obtained by plate culture.

Almost every species detected by plate culture was not found by PCR-DGGE, while,

PCR-DGGE detected a diversity of bacterial species that were not recovered by plate culture (Table 3.6, Fig. 3.13). For example, L. plantarum found by plate culture of both

Shiraz and Semillon grape samples at stage I, was not detected by PCR-DGGE. It was also detected at this stage by enrichment cultures of the Shiraz grapes. The diversity of species found by PCR-DGGE, did not suggest any obvious ecological trends.

G G 85

Table 3.6 Bacterial species and populations (CFU/g) on grapes at different stages of maturity in vineyard C during the 2002-2003 season as determined by (a) plate culture, (b) PCR-DGGE and (c) enrichment culture

Grape Detection Maturity of grapes variety method I II III IV V a B. thuringiensis (3.2u105) B. thuringiensis (1.5u103) B. thuringiensis (4.5u102) B. thuringiensis (1.0u102) B. thuringiensis (1.0u102) Lb. plantarum (5.0u102) Pa. agglomerans (2.5u102) b B. thuringiensis B. thuringiensis B. thuringiensis Ac. radioresistens B. thuringiensis Acinetobacter jonhsonii Ar. gandensis G. tepidamans Er. herbicola B. benzoevorans Ralstonia sp. B. drentensis B. sphaericus Shiraz Bacillus sp. G. tepidamans Microbacterium sp. Ralstonia sp. S. proteamaculans c B. thuringiensis B. thuringiensis B. thuringiensis B. thuringiensis B. thuringiensis Lb. plantarum B. mycoides a B. thuringiensis (2.4u105) B. thuringiensis (7.8u103) B. thuringiensis (1.0u103) B. thuringiensis (5.0u101) B. thuringiensis (5u101) Lb. plantarum (6.0u102) S. marcescens (7.5u102) C. flaccumfaciens (1.0u102) B. megaterium (1.5u102) b B. thuringiensis B. thuringiensis B. thuringiensis Mb. imperiale Ralstonia sp. Bradyrhizobium sp. Erwinia sp. Methylobacterium sp. Sphingomonas sp. Semillon Friedmanniella sp. Ralstonia sp. Ps. putida Strc. thermophilus Pantoea sp. Sphin. terrae Streptomyces sp. Uncultured bacterium Staph. saprophyticus clone Uncultured bacterium gene c B. thuringiensis B. thuringiensis B. thuringiensis B. thuringiensis B. thuringiensis

G 86

Shiraz Semillon M I II III IV V M I II III IV V

B Staph s Ac r

Be Ps p Ac j Bt Bd St th Bt Sphin * B Pa Sp Vv Vv Meth Sphin t Gt Rals Rals Brad M Mim Rals* Un Fried Er h Str Ar g Un Er

Fig. 3.13 PCR-DGGE analysis of bacterial DNA in rinses of wine grapes from vineyard C at different stages during cultivation in 2002-2003. Lanes: I, II, III, IV, V, represent the stages of maturity referred to in Table 3.2. M, marker referred to in Fig. 3.5. Ac j, Acinetobacter johnsonii; Ac r, Acinetobater radioresistens; Ar g, Ar. gandensis; B, Bacillus sp.; Bd, B. drentensis; Be, B. benzoevorans; Bt, B. thuringiensis; Brad, Bradyrhizobium sp.; Er, Erwinia sp.; Er h, Er. herbicola; Fried, Friedmanniella sp.; Gt, G. tepidamans; M , Microbacterium sp.; Mim, Mb. imperiale; Meth, Methylobacterium sp.; Pa, Pantoea sp.; Ps p, Ps. putida; Rals, Ralstonia sp.; Sp, S. proteamaculans; Sphin, Sphingomonas sp.; Sphin t, Sphin. terrae; Str th, Strc. thermophilus; Str, Streptomyces sp.; Staph s, Staph. saprophyticus; Un, Unidentified bacterial isolates; Vv, Vitis vinifera.*, plant DNA; Vv, Vitis vinifera. G 87

3.3.3.3 Mudgee region, 2002-2003

Vineyards D and E, located in the Mudgee region, were separated by a distance of about

5 kilometres. Vineyard D was a relatively new plantation, about five years old, whereas vineyard E was an older, more well established vineyard (about 20 years). Both vineyards were subjected to the same protocol of agrichemical applications.

Bacterial populations on grapes from these vineyards were low at about 101-103 CFU/g, except for some higher initial populations of B. thuringiensis on the Shiraz and Merlot varieties (104-106 CFU/g, maturity stage I). Application of Bt to grapes in vineyards D and E was done only until veraison. Thus, B. thuringiensis on grapes in these vineyards was only detected by plate culture or PCR-DGGE at the earlier stages (Tables 3.7, 3.8).

However, its presence on grapes throughout all stages of cultivation was detected by enrichment culture. Plate culture gave a diversity of bacterial isolates with little consistency in their recovery. Curtobacterium flaccumfaciens was detected at 102-103

CFU/g on all varieties at harvest or near harvest (maturities IV-V). It was also found at earlier stages of maturation. Species of Arthrobacter were also frequently detected by plate culture. PCR-DGGE analysis gave a different spectrum of isolates. Most notable was the consistent detection of B. benzoevorans from the Shiraz, Chardonnay and

Cabernet Sauvignon grapes (Fig. 3.14).

Lactic acid bacteria were not isolated from grape samples by plate culture and PCR-

DGGE. There were three occasions where E. mundtii was isolated by enrichment culture, but not by plate culture or PCR-DGGE, from Chardonnay (maturity stages III and IV) and Merlot (maturity stage V).

G G 88

Table 3.7 Bacterial species and populations (CFU/g) on grapes at different stages of maturity in vineyard D during the 2002-2003 season as determined by (a) plate culture, (b) PCR-DGGE and (c) enrichment culture

Grape Detection Maturity of grapes variety method I II III IV V Shiraz a B. thuringiensis (2.6u104) B. thuringiensis (5.0u101) B. thuringiensis (5.0u101) C. flaccumfaciens (1.0u102) Pa. ananatis (5.0u102) Arthrobacter sp. (3.0u102) Arthrobacter sp. (1.5u102) Sp. parapaucimobilis Pa. agglomerans (2.0u102) Oxalobacteraceae bacterium (5.0u101) Rathayibacter tritici (4.5u102) (2.0u102) Pa. agglomerans (2.5u102) b B. thuringiensis B. benzoevorans B. benzoevorans B. benzoevorans G. tepidamans B. benzoevorans C. flaccumfaciens c B. thuringiensis B. thuringiensis B. thuringiensis B. thuringiensis B. thuringiensis Pa. agglomerans Er. rhaphontici B. mycoides Pseudomonas sp. B. sphaericus Chardonnay a B. thuringiensis (5.0u101) B. thuringiensis (5.0u101) Ar. gandensis (3.5u102) Arthrobacter sp. (5.0u101) Arthrobacter sp. Ar. polychromogenes (4.5u102) Rathayibacter tritici Arthrobacter sp. (4.0u102) B. fumarioli (5.0u101) (5.0u101) Curtobacterium sp. (3.5u102) (5.0u102) C. flaccumfaciens (3.0u102) C. fangii (1.0u102) Microbacterium sp. (7.5u102) Frigoribacterium sp. (3.5u102) C. flaccumfaciens Mb. terregens (7.5u102) Rathayibacter tritici (3.5u102) (2.0u102) Rh. equi (3.0u102) b B. benzoevorans B. benzoevorans B. benzoevorans B. benzoevorans B. benzoevorans Staphylococcus sp. c B. thuringiensis B. thuringiensis B. thuringiensis B. thuringiensis B. thuringiensis Pa. ananatis Br acetylicum E. mundtii E. mundtii Pa. agglomerans

G G 89

Table 3.8 Bacterial species and populations (CFU/g) on grapes at different stages of maturity in vineyard E during the 2002-2003 season as determined by (a) plate culture, (b) PCR-DGGE and (c) enrichment culture

Grape Detection Maturity of grapes variety methods II III IV V Cabernet a B. thuringiensis (1.0u102) B.thuringiensis (1.5u102) B. megaterium (1.0u102) Microbacterium sp. (5u101) C. flaccumfaciens (2.0u102) Sauvignon B. megaterium (4.5u102) Arthrobacter sp. (1.0u102) Rhodococcus sp. (5u101) C. flaccumfaciens (3.0u102) Rhodococcus sp. (2.5u102) Microbacterium sp. (4.0u102) Mb. phyllosphaerae.(4.0u102) Rh. equi (2.0u102) b B. benzoevorans B. benzoevorans B. benzoevorans B. circulans B. benzoevorans Ralstonia sp. Streptomyces sp. Mycobacterium sp. c B. thuringiensis B. thuringiensis B. thuringiensis B. thuringiensis B. thuringiensis Acinetobacter sp. Pa. agglomerans Pa. stewartii Merlot a B. thuringiensis (1.1u106) B.thuringiensis (5.0u101) Arthrobacter sp. (1.0u102) ND C. flaccumfaciens (1.0u102) Microbacterium sp. (6.0u102) Pa. agglomerans (5.0u101) Microbacterium sp. (3.5u102) Arthrobacter sp. (5.0u101) Rhodococcus sp. (1.0u102) Mb. paraoxydans (1.0u102) b B. thuringiensis B. maroccanus Acinetobacter sp. Uncultured bacterium Escherichia coli B. megaterium Nostoc sp. Rhodococcus sp. Frankiaceae sp. Uncultured rhizosphere bacterium c B. thuringiensis B. thuringiensis B. thuringiensis B. thuringiensis B. thuringiensis Pantoea sp. Pa. agglomerans Br. acetylicum E. mundtii Pseudomonas sp. B. megaterium

G G 90

Table 3.8 (continued) Bacterial species and populations (CFU/g) on grapes at different stages of maturity in vineyard E during the 2002- 2003 season as determined by (a) plate culture, (b) PCR-DGGE and (c) enrichment culture

Grape Detection Maturity of grapes variety methods II III IV V Sauvignon a B. thuringiensis (1.7u103) B. thuringiensis (1.0u102) Sphingomonas sp. (3.0u102) C. flaccumfaciens (4u102) B. pumilus (3.5u102) Blanc C. flaccumfaciens (5.0u102) Arthrobacter sp. (5.0u101) C. flaccumfaciens (5.0u101) Frigoribacterium sp. (1.5u103) Mb. barkeri (5.0u101) Mb. paraoxydans (6.8u102) b B. thuringiensis ND Uncultured bacterium clone Pl. psychrotoleratus Pseudoalteromonas spongia Rhodococcus sp. Strm. scabiei Uncultured Actinobacteridae Mycobacterium tuscia Uncultured earthworm cast Methylobacterium sp. bacterium clone Tsukamurella tyrosinosolvens Mycobacterium tuscia c B. thuringiensis B. thuringiensis B. thuringiensis B. thuringiensis B. thuringiensis Acinetobacter sp. Pa. ananatis Pa. ananatis. Pseudomonas sp. ND, Not detected

G Vineyard D Vineyard E

Shiraz Chardonnay Cabernet Sauvignon Merlot Sauvignon Blanc M I II III IV V M I II III IV V I II III IV V M I II III IV V M I II III IV V

Acin Bm Ps sp Staph Be Bc Bma Be Bt Be Nos Bt Ec Bt Cf Vv Rals Un Vv Vv Gt Vv Meth Fran Pl p Un Strem Un Strem s Rhodo Tt Un UnRhodo Myco Myco

Fig. 3.14 PCR-DGGE analysis of microbial DNA in rinses of wine grapes sample from vineyards D and E at different stages during cultivation in 2002-2003. Lanes: I, II, III, IV, V, stages of maturity referred to in Table 3.2. M, marker referred to in Fig. 3.5. Acin, Acinetobacter sp.; Bc, B. circulans; Be, B. benzoevornas; Bm, B. megaterium; Bma, B. marococcanus; Bt, B. thuringiensis; Cf, C. flaccumfaciens; Ec, E. coli; Fran, Frankiaceae sp.; Gt, G. tepidamans; Meth, Methylobacterium sp.; Myco, Mycobacterium sp.; Nos, Nostoc sp.; Pl p, Pl. psychrotoleratus; Ps sp, Pseudoalteromonas spongia; Rals, Ralstonia sp.; Rhodo, Rhodococcus sp.; Staph, Staphylococcus sp.; Strem, Streptomyces sp.; Strem s, Strm. scabiei; Tt, Tsukamurella tyrosinosolven; Un, Uncultured isolates; Vv, Vitis vinifera. 91 G 92

3.3.3.4 Griffith region, 2002-2003

Table 3.9 shows the bacterial species and populations detected on Shiraz and

Chardonnay grapes harvested from vineyard F. Bacterial populations on grapes were relatively low throughout cultivation, at approximately 102-103 CFU/g, except for higher populations of B. thuringiensis (104-105 CFU/g) at maturity stage I on both grape varieties (Table 3.9). Bacillus thuringiensis was mostly found on samples of Shiraz grapes by agar culture but not on Chardonnay grapes by this method except at maturity stage I. However, its presence was consistently detected by enrichment culture. As determined by plate culture, C. flaccumfaciens (102-104 CFU/g) was the most prevalent species on Chardonnay grapes at most stages of maturation. Species of Arthrobacter

(102-103 CFU/g) were also frequently isolated. Neither of these species were detected by PCR-DGGE, but PCR-DGGE revealed the consistent presence of B. benzoevorans which was not detected by plate and enrichment cultures.

Lactic acid bacteria were not isolated from grape samples by plate culture. Enterococcus mundtii was isolated from Chardonnay grapes (maturity stages III-V) by enrichment culture and Lc. lactis was detected on Shiraz grapes (maturity stage III) by PCR-DGGE but not by enrichment culture (Fig. 3.15). The diversity of Bacillus species (B. megaterium, B. senegalensis, B. sphaericus) on grapes from this vineyard is notable.

G G 93

Table 3.9 Bacterial species and populations (CFU/g) on grapes at different stages of maturity in vineyard F during the 2002-2003 season as determined by (a) plate culture, (b) PCR-DGGE and (c) enrichment culture

Grape Detection Maturity of grapes variety method I II III IV V Shiraz a B. thuringiensis (4.1u104) B. thuringiensis (5.0u101) B. thuringiensis (5.0u101) Arthrobacter sp. (1.0u102) B. thuringiensis (5.0u101) Arthrobacter sp. (1.5u102) B. megaterium (1.0u101) B. megaterium (5.0u101) B. senegalensis (5.0u101) Pa. agglomerans (2.0u102) Leifsonia poae (1.0u101) Ar. polychromogenes (5.0u101) Mc. luteus (5.0u101) b B. thuringiensis B. benzoevorans B. benzoevorans B. benzoevorans B. benzoevorans Lc. lactis c B. thuringiensis B. thuringiensis B. thuringiensis B. thuringiensis B. thuringiensis B. megaterium Pa. agglomerans Chardonnay a B. thuringiensis (1.3u105) B. thuringiensis (5.0u101) C. flaccumfaciens Ar. oxydans (9.0u102) Ar. polychromogens. Ar. oxydans (1.5u102) (1.7u103) B. megaterium (2.0u102) (3.0u102) C. flaccumfaciens (5.0u103) B. senegalensis (2.0u102) C. flaccumfaciens (5.5u102) Pa. toletana (1.4u103) C. flaccumfaciens (4.9u103) Microbacterium sp. (1.0u102) b B. thuringiensis B. benzoevorans B. benzoevorans B. benzoevorans B. benzoevorans Mb. imperiale c B. thuringiensis B. thuringiensis B. thuringiensis B. thuringiensis B. thuringiensis B. megaterium E. mundtii E. mundtii B. sphaericus Pseudomonas sp. E. mundtii Br. acetylicum G

G 94

Shiraz Chardonnay MIIIIIIIVV MI IIIIIIVV

Be

Bt Be Lc l

Bt Vv

Mim Vv

Fig. 3.15 PCR-DGGE analysis of bacterial DNA in rinses of wine grapes from vineyard F at different stages during cultivation in 2002-2003. Lanes: I, II, III, IV, V, represent the stages of maturity, referred to in Table 3.2. M, marker referred to in Fig. 3.5. Be, B. benzoevornas; Bt, B. thuringiensis, Mim, Mb. imperiale; Lc l. Lc. lactis; Vv, Vitis vinifera.

3.3.4 Bacterial species associated with undamaged and damaged wine

grapes

Comparisons of total bacterial populations on undamaged and damaged grapes have

been presented in Section 3.2.2. For red grape varieties, there were little differences

between total populations on undamaged and damaged grapes while, for white varieties,

damaged grapes tended to have slightly higher populations than undamaged grapes.

These trends were generally reflected in data for individual species (Tables 3.10, 3.11).

Bacillus thuringiensis and C. flaccumfaciens, to a lesser extent, were the most

frequently cultured species from both undamaged and damaged grapes. Generally, their

populations were quite low, at 102 -103 CFU/g, but C. flaccumfaciens occurred on G 95 damaged Chardonnay grapes, vineyard A at 2.6u104 CFU/g. On three occasions,

Gluconobacter oxydans (Shiraz, vineyard A, Chardonnay, vineyard A, Semillon, vineyard A) was isolated from damaged grapes, but not undamaged grapes, at 103-105

CFU/g. An unidentified species of Pantoea was significant (103-105 CFU/g) on undamaged and damaged Semillon grapes from vineyard A. Lactic acid bacteria were cultured from the grapes on only three occasions, and these were Enterococcus species at low populations (101-102 CFU/g) from damaged Shiraz and Chardonnay grapes.

In most cases, species found by culture on agar plates were also detected by PCR-

DGGE. Some exceptions were C. flaccumfaciens and Pantoea spp. on damaged

Chardonnay and Semillon from vineyard A. Even though their populations were shown at 104 CFU/g by plate culture, they were not detected by PCR-DGGE. Generally, bacterial species with populations of less than 102 CFU/g, were hardly detected by

PCR-DGGE.

Nevertheless, PCR-DGGE revealed the presence of a greater diversity of species than plate culture. For example, Lactobacillus kunkeei, Acetobacter diazotrophicus and species of Gluconobacter, in addition to G. oxydans were detected by PCR-DGGE but not plate culture on damaged grapes of Cabernet Sauvignon and Semillon from vineyard A. Bacillus benzoevorans was detected on Shiraz (vineyards D, F), Cabernet

Sauvignon (vineyard E) and Chardonnay (vineyards D, F) grapes by PCR-DGGE, but not by plate culture.

G G 96

Table 3.10 Bacterial species on undamaged and damaged red grapes at commercial harvest time from several vineyards as determined by (a) plate culture and (b) PCR- DGGE

Grape Detection Vineyard Undamaged Damaged variety method B. thuringiensis (5.5u102) B. thuringiensis (3.6u102) a C. flaccumfaciens (1.9u103) G. oxydans (1.4u103) A B. thuringiensis C. flaccumfaciens G. oxydans b C. flaccumfaciens Br. equis Uncultured bacterium clone Mb. imperiale a B. thuringiensis (2.0u102) B. thuringiensis (4.0u102) A b B. thuringiensis B. thuringiensis a Pseudomonas sp. (5.0u101) B. megaterium (5.0u101) B Lc. lactis Lc. lactis b Endosymbiont of Brevipalpus Endosymbiont of Brevipalpus lewisi lewisi B. thuringiensis (1.0u102) B. thuringiensis (1.0u102) a 2 Shiraz C. flaccumfaciens (1.0u10 ) B. thuringiensis Hafnia alvei C Ge. tepidamans Ge.tepidamans b B. subtilis Ralstonia sp. Uncultured bacterium clone Pa. ananatis (5.0u102) E. durans (1.0u102) a 1 D Sp.parapaucimobilis (5.0u10 ) B. benzoevorans ND b Ge. tepidamans B. thuringiensis (5.0u101) B. megaterium (1.5u102) B. senegalensis (5.0u101) C. flaccumfaciens (5.0u101) a 1 F Arthrobacter sp. (5.0u10 ) Mc. luteus (5.0u101) b B. benzoevorans B. benzoevorans B. thuringiensis (1.3u103) B. thuringiensis (1.2u103) a C. flaccumfaciens (2.5u103) C. flaccumfaciens (7.5u102) Erwinia sp. (6.1u103) A B. thuringiensis Lb. kunkeei b C. flaccumfaciens A. diazotrophicus Pseudomonas sp. G. cerinus Cabernet Sphingomonas sp. G. oxydans Sauvignon a B. thuringiensis (2.0u102) B. thuringiensis (7.5u102) A Sphingomonas sp. B. thuringiensis b Lc. lactis Uncultured bacterium clone a C. flaccumfaciens (2.0u102) Erwinia rhapontici (1.0u102) E b B. benzoevorans ND a C. flaccumfaciens (1.0u102) B. thuringiensis (1.5u102) Merlot E b Escherichia coli ND ND, Not detected

G G 97

Table 3.11 Bacterial species on undamaged and damaged white grapes at commercial harvest time from several vineyards as determined by (a) plate culture and (b) PCR- DGGE

Grape Detection Vineyard Undamaged Damaged variety method B. thuringiensis (1.3u103) B. thuringiensis (1.0u103) a C. flaccumfaciens (3.4u103) C. flaccumfaciens (2.6u104) Pseudomonas sp. (1.4u103) G. oxydans (5.0u104) A B. thuringiensis Lb. kunkeei Sphingomonas sp. A. diazotrophicus b C. flaccumfaciens G. cerinus G. oxydans B. thuringiensis (4.5u102) B. thuringiensis (1.2u103) C. flaccumfaciens (1.5 102) a u A Arthrobacter sp. (1.0u102) Pseudomonas sp. (4.5u102) b B. thuringiensis B. thuringiensis Chardonnay a ND ND B Lc. lactis Lc. lactis b Endosymbiont of Brevipalpus Endosymbiont of Brevipalpus lewisi lewisi C. flaccumfaciens (2.0u102) C. flaccumfaciens (2.0u102) a Arthrobacter sp. (5.0X101) Arthrobacter sp. (5.0u101) D C. fangii (1.0u102) E. mundtii (1.0u102) b B. benzoevorans B. benzoevorans C. flaccumfaciens (4.0u102) C. flaccumfaciens (5.5u102) Ar. polychromogens (3.0 102) B. megaterium (2.0 102) a u u F B. megaterium (1.5u102) E. mundtii (1.0u101) Microbacterium sp. (1.0u102) b B. benzoevorans B. benzoevorans B. thuringiensis (2.0u102) B. thuringiensis (5.5u102) a C. flaccumfaciens (1.0u103) Pantoea sp. (6.7u104) Pantoea sp. (2.0u103) G. oxydans (4.0u105) A B. thuringiensis A. diazotrophicus C. flaccumfaciens G. cerinus b Ps. putida G. oxydans Pantoea sp. Semillon Sphingomonas sp. a B. thuringiensis (5.5u102) B. thuringiensis (2.6u102) A b B. thuringiensis Uncultured bacterial gene a B. thuringiensis (5.0u101) ND Strc. thermophilus ND C Sphingomonas sp. b Ralstonia sp. Uncultured bacterium clone (Un) a B. pumilus (3.5u102) Pantoea sp. (7.9u102) Pseudoalteromonas spongia ND Sauvignon E Uncultured actinomycetes Blanc b Methylobacterium sp. Tsukamurella tyrosinosolvens Mycobacterium sp. ND, not detected

G Vineyard A, 2001-2002 Vineyard A, 2002-2003 Vineyard E, 2002-2003

Shiraz Chardonnay Shiraz Cabernet Sauvignon Merlot MUD MUD MUMD D MUD IV UDM Lb k

Cf Lc l Cf Bt Bt Un Bt Sphin Bt Cf Ec Vv Sphin Vv Br e Cf Vv Vv

Mim A diaz G oxy G cer A diaz G oxy

Fig. 3.16 PCR-DGGE analysis of bacterial DNA in rinses of undamaged (U) and damaged (D) wine grapes from several vineyards at commercial harvest time. Lanes: V, represents the stage of maturity at harvest; D, damaged grapes at maturity stage V; M, marker referred to in Fig. 3.5. A diaz, A. diazotrophicus; Bt, B. thuringiensis; Br e, Br. equis; Cf, C. flaccumfaciens; Ec, E.coli; G cer, G. cerinus; G oxy, G. oxydans; Lb k, Lb. kunkeei; Lc l, Lc. lactis; Mim, Mb. imperiale; Sphin, Sphinogomonas sp.; Un, Uncultured bacterium clone; Vv, Vitis vinifera. 98 G 99

3.3.5 Frequency of isolation of bacteria from wine grape samples

Bacillus thuringiensis was the most frequently isolated bacterial species from red and white wine grapes, as determined by all three methods, plate culture, enrichment culture and PCR-DGGE (Tables 3.12, 3.13). However, it was much less frequently detected by

PCR-DGGE, than by culture. Curtobacterium flaccumfaciens was the next most frequently isolated species by plate culture, followed by Pseudomonas and Arthrobacter spp. A diversity of other species was isolated by plate culture, but these were infrequent and sporadic. After B. thuringiensis, PCR-DGGE analysis showed B. benzoevorans, to be the most frequently detected species. This species was never detected by culture methods. PCR-DGGE analysis also confirmed the prevalence of C. flaccumfaciens on grapes. After these few species, there is little agreement between the phyllospheric microflora revealed by plate culture and PCR-DGGE. Numerous species detected by plate culture were not detected by PCR-DGGE. Numerous species detected by PCR-

DGGE were not found by plate culture. Of the three analytical methods, enrichment culture revealed the least biodiversity showing, on most occasions, only the presence of

B. thuringiensis (Table 3.12)

G G 100

Table 3.12 Main bacterial species isolated from undamaged grapes at the time of commercial harvest from all vineyards during the 2001-2002 and 2002-2003 seasons, as determined by plate culture and enrichment culture methods

Grape variety Red White Red White No. of samples 20 16 18 14 Detection method Plating Enrichment Bacillus thuringiensis 11a 9 16 14 Curtobacterium flaccumfaciens 4 7 0 0 Arthrobacter sp. 1 3 0 0 Pseudomonas sp. 1 3 0 0 Bacillus megaterium 0 2 0 0 Bacillus sphaericus 0 0 0 2 Pantoea ananatis 2 0 0 0 Enterococcus mundtii 0 0 1 1 Sphingomonas sp. 1 0 0 0 Bacillus pumilus 0 1 0 0 Bacillus senegalensis 1 0 0 0 Curtobacterium fangii 0 1 0 0 Micrococcus luteus 1 0 0 0 Pantoea sp. 0 1 0 0 Total 22 27 17 17 a, Number of times isolated

Table 3.13 Main bacterial species isolated from undamaged grapes at the time of commercial harvest time from all vineyards during the 2001-2002 and 2002-2003 seasons, determined by PCR-DGGE

Grape variety Red White No. of samples 20 16 Bacillus thuringiensis 5a 5 Bacillus benzoevorans 5 4 Curtobacterium flaccumfaciens 2 4 Sphingomonas sp. 2 3 Endosymbiont of Brevipalpus lewisi 2 2 Geobacillus tepidamans 2 0 Lactococcus lactis 0 2 Pseudomonas sp. 1 0 Brevibacterium equis 1 0 Escherichia coli 1 0 Methylobacterium sp. 0 1 Microbacterium imperiale 1 0 Microbacterium sp. 0 0 Mycobacterium sp. 0 1 Pantoea sp. 0 1 Pseudoalteromonas spongia 0 1 Pseudomonas putida 0 1 Ralstonia sp. 0 1 Total 22 26 a, Number of times isolated

G G 101

3.4 DISCUSSION

This Chapter reports the first systematic investigation of bacteria associated with wine grapes harvested from Australian vineyards. To maximise the validity and utility of the conclusions, grapes representing six different varieties and harvested from six different vineyards were examined. Grapes from one vineyard were examined over two ,

2001-2002 and 2002-2003. Three analytical methods were used - plate culture, enrichment culture and PCR-DGGE.

3.4.1 Population and species diversity

The total bacterial populations on healthy, undamaged grapes at the time of commercial harvest were consistently low at 102-103 CFU/g. Similar conclusions were reported in studies of wine grapes in other countries (Lafon-Lafourcade et al. 1983; Manca de

Nadra and Strasser de Saad 1987; Sieiro et al. 1990; de la Torre et al. 1998). Higher populations were found on grapes at earlier stages of cultivation, and this correlated with application of the bioinsecticide Bt, and recovery of B. thuringiensis on isolation media.

Bacillus thuringiensis was the most prevalent bacterial species on wine grapes, and was isolated from almost every sample taken throughout cultivation. Its discovery and predominance was unexpected because it has not been mentioned in previous literature on wine microbiology (Kunkee 1967; Fleet 1993; Fugelsang 1997; Ribéreau-Gayon et al. 2000). However, after reviewing viticultural practices in Australian vineyards, its discovery could be predicted and correlated with the widespread application of biological pesticide preparations (e.g. BT, Dipel Forte, Delfin WG, Success) to control

G G 102 apple moth and other insect infestations (Emmett et al. 1992). These bioinsecticides are essentially powdered or granulated preparations of the bacterium, B. thuringiensis, that are reconstituted in water and sprayed onto vines and grape berries. Generally, the bioinsecticide is applied at regular intervals from bud burst until 1-2 weeks before grape harvest, and this explains the consistent isolation of B. thuringiensis from wine grapes throughout cultivation. B. thuringiensis is an endospore forming organism, consequently, it is not unexpected to isolate it from grapes at harvest, when it would be several days or weeks after the last time it was sprayed onto the vines. Interestingly, B. thuringiensis was recovered from the grapes in vineyard B which is an organic vineyard, that had not been exposed to agrichemicals or pesticide applications for many years. In this case, it was not recovered by plate culture or PCR-DGGE methods, but only after enrichment culture, suggesting a very low level of presence. Presumably, spores of the organism had survived in the environment for many years, or arrived on the grapes by soil or dust contamination from other nearby vineyards. Bacillus thuringiensis has been widely used as a bioinsecticide in agricultural practices, for over 50 years (Glare and O’Callaghan

2000). Some years ago, Bidochka et al. (1987) briefly reported its isolation from table grapes and other fruits, and linked its presence to biopesticide applications .The prevalence of B. thuringiensis on wine grapes could have significant implications in the process of wine making, and this topic is examined and discussed in more detail in

Chapter 5.

After B. thuringiensis, C. flaccumfaciens was the next most prevalent bacterial species detected on wine grapes. Its populations were low and rarely exceeded 103-104 CFU/g.

It is one of the most abundant epiphytic bacterial species found on the surface of plants

(i.e. sugar beet, mango, peanut) (Thompson et al. 1993; Sundin and Jacobs 1999; de

G G 103

Jager et al. 2001). It is a chromogenic, pigmented bacterium that produces yellow to orange coloured colonies on agar plating media (Sundin and Jacobs 1999; Jacobs and

Sundin 2001), making it easy to detect and recognise. It was clearly seen after culture of grape rinses on PCA and readily identified by ribosomal DNA sequencing. However, strain variation within the species seems likely, as colonies with slightly different pigmentation properties were observed, but sequenced as C. flaccumfaciens.

Pigmentation is believed to confer UV radiation resistance to microorganisms (Lindow

1991; Hirano and Upper 2000), and this property may explain its prevalence on the grape surface. The ability to produce biosurfactants, and utilize or interface with grape cutin or epicuticular waxes are other properties that could select for the occurrence of C. flaccumfaciens at the grape surface, and these properties need further research. Previous studies have reported the isolation of C. flaccumfaciens from the xylem of grape vines

(Bell et al. 1995) and from grape ice musts (Subden et al. 2003). Curtobacterium flaccumfaciens has not been considered in previous literature as an organism of significance in wine production. It is not known if its presence might influence the colonization of grapes by filamentous fungi, yeasts, lactic acid bacteria, or acetic acid bacteria and these questions require further research. Other bacteria, less frequently and sporadically found on wine grapes were species of Arthrobacter, Bacillus,

Microbacterium, Pantoea, Pseudomonas, and Sphingomonas.

Bacillus benzoevorans was never isolated from grapes by cultural methods, but it was consistently detected by culture-independent PCR-DGGE analysis of grapes throughout cultivation in several vineyards. Its broad distribution (Hunter Valley, Mudgee, Griffith regions) and persistent detection throughout cultivation suggest that B. benzoevorans may be a significant species and have an important role in the ecosystem of wine grapes.

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It cannot be concluded from the data obtained, if it occurs as dead, non-viable cells or as viable but not culturable cells. Nevertheless, there is clear evidence of its presence, and its significance needs to be considered in future research. This species has not been reported on wine grapes before, but it is associated with soil ecosystems where it was more frequently detected by culture-independent, molecular methods than by direct culture (Felske et al. 1998; Garbeva et al. 2003; Tzeneva et al. 2004). It has been isolated from feed lot manure, as well (Klieve et al. 1999). With this ecology, it is not unexpected that it might be associated with grapes that are constantly exposed to dust and soil.

Of the 93 samples of healthy undamaged grapes that were examined from all maturity stages, only 15 showed the presence of lactic acid bacteria, and no acetic acid bacteria were detected, by any method. Based on literature reports (Wibowo et al. 1985; Fleet

1993; Fugelsang 1997), a greater incidence of these bacteria was expected. Nevertheless, several other studies have also noted the difficulty and inconsistency in finding these bacteria on wine grapes (Lafon-Lafourcade et al. 1983; Mance de Nadra et al. 1987;

Sieiro et al. 1990; de la Torre et al. 1998). Possibly, these bacteria are less prevalent on healthy, undamaged grapes than previously thought. Of the lactic acid bacteria, only Lc. lactis, Lb. plantarum, E. faecalis and E. mundtii, were found in this study.

Lactobacillus plantarum has been found on wine grapes in other studies (Weiller and

Radler 1970; Lafon-Lafourcade et al. 1983; Edinger and Splittstoesser 1986; Suárez et al. 1994), but not Lc. lactis or Enterococcus species. These latter species have not been reported to be significant or active in wine fermentation. The failure to find the principal malolactic organism, O. oeni, on grapes is notable but not unexpected as many previous

G G 105 researchers have not detected its presence on various samples of wine grapes (Sieiro et al. 1990; de la Torre et al. 1998). It appears that this important species has never been isolated from wine grapes, but its isolation was reported from the leaves of wine grapes

(Weiller and Radler 1970). More detailed investigation of lactic acid bacteria on wine grapes using enrichment cultures is presented in Chapter 4.

There are only a few reports on the isolation of acetic acid bacteria (Gluconobacter,

Acetobacter) from wine grapes (Joyeux et al. 1984; Lafon-Lafourcade and Ribéreau-

Gayon 1984; Drysdale and Fleet 1988; du Toit and Pretorius 2000), and it is possible that these findings reflect more on their isolation from damaged berries than undamaged berries (see later). Specific media for the isolation of these bacteria such as WL Nutrient agar and glucose yeast extract carbonate agar (Drysdale and Fleet 1988) were not used in this study, and it is a limitation of the analytical protocol used. However, these bacteria grow well on MRS based media (Hommel and Ahnert 2000) as used in this study. They were not detected by PCR-DGGE analysis of rinses from non damaged grapes, but this method did detect them on damaged grapes (see later).

3.4.2 Analytical strategies for monitoring bacterial species associated

with wine grapes

Three analytical approaches were used to investigate the bacterial ecology of grapes- plate culture to detect and quantify populations (detection limit 10 CFU/mL of rinse), enrichment culture to detect species at very low populations, and PCR-DGGE as a culture-independent method that might recover viable but non culturable species. The predominance of B. thuringiensis in the grapes was demonstrated by all three methods.

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The next most prevalent species, C. flaccumfaciens was confirmed by both plate culture and PCR-DGGE methods. Thereafter, the three methods gave diverse data.

Enrichment culture revealed the least species diversity, with B. thuringiensis dominating in almost every analysis (Table 3.13). This finding suggests the greater presence of cells or spores of this organism on grapes and their ability to grow quickly and dominate in the enrichment cultures - Nutrient broth and MRS broth. Even though the MRS medium is designed to encourage the growth of lactic acid bacteria, there were only nine occasions where lactic acid bacteria were found (e.g. E. mundtii (7), Lb. plantarum (1),

Lc. lactis (1)). There were numerous grape batches from which no lactic acid bacteria could be cultured. Possibly, the growth of B. thuringiensis in this medium could have suppressed the growth and detection of lactic acid bacteria, and these issues are addressed in more detail in Chapter 4.

Bacillus thuringiensis and C. flaccumfaciens were readily detected by both plate culture and PCR-DGGE methods. Their colony morphologies were quite distinct on agar media.

The banding pattern of B. thuringiensis on gels after DGGE was very characteristic, one major band plus three weaker DNA bands (Fig. 3.5), suggesting that the organism has multiple copies of the 16S rRNA gene that was amplified by PCR (Farrelly et al. 1995;

Nübel et al. 1996). Although pure cultures of C. flaccumfaciens gave one dominant band (Fig. 3.5), some isolates from grapes occasionally gave two bands that sequenced as C. flaccumfaciens (e.g. Shiraz, Semillon, Vineyard A, 2001-2002, Fig. 3.10). This finding could reflect the presence of more than one strain of this species on grapes.

Such possibility is also suggested and supported by the heterogeneity in colony morphology of this organism on agar culture media, as mentioned previously.

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After B. thuringiensis and C. flaccumfaciens, the bacterial flora of grapes as detected by either plate culture or PCR-DGGE were somewhat different. Species detected by one method were not detected by the other method (Tables 3.12 - 3.13). The most obvious differences were the detection of B. benzoevorans and various Gram negative bacteria, particularly, species of Pseudomonas, Sphingomonas and Ralstonia by PCR-DGGE but not by plate culture. As mentioned already (Chapter 2), the surface of grapes represents an adverse phyllospheric habitat which is subject to significant fluctuation of temperature, UV irradiation and water availability, and is low in nutrient availability.

Spore-forming bacteria (Bacillus spp.) would be favoured by such conditions, but other bacterial species could be induced into the VBNC state. In fact, species of Pseudomonas

Sphingomonas and Ralstonia are frequently reported as one of dominant colonizers on the surface of plants and as species that readily enter the VBNC state (Lindow and

Brandl 2003). The concept of VBNC organisms in grape and wine ecosystems is not widely recognised, but has been reported by Millet and Lonvaud-Funel (2000) for lactic acid bacteria and acetic acid bacteria in wines. Culture-independent, molecular methods such as PCR-DGGE are needed to detect the presence of VBNC cells.

3.4.3 Factors affecting the bacterial ecology of wine grapes

As discussed in Fleet et al. (2002) and Chapter 2 of this thesis, several intrinsic and extrinsic factors have the potential to influence the occurrence of microorganisms on wine grapes. These include grape cultivar, climate effects, region, application of agrichemicals and berry damage. A systematic, controlled investigation of these variables was beyond the time frame and scope of this study, but tentative observations and conclusions can be advanced.

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The surface properties of grapes are determined by the presence of cutin and epicuticular waxes, the contents and composition of which vary to some extent with grape variety (Hill et al. 1981; Hardie et al. 1996; Kök and ÇeĒk 2004). Such variations could affect the surface microflora. Although it has been reported that grape cultivar could influence the diversity of yeasts associated with the grape surface (de la Torre et al. 1999; Sabate et al. 2002), similar populations and bacterial species were found on all the grape varieties examined in this study. No correlation was observed between the presence of any particular bacterial species and grape variety for healthy, undamaged berries.

Only one vineyard, vineyard A was examined over two vintages for bacterial flora. For this vineyard, the 2001-2002 vintage was characterised by slightly cooler temperature and more rainfall than the 2002-2003 vintage. It is generally reported that grapes from higher rainfall vintages have higher, and more diverse microbial populations (de la

Torre et al. 1998). This trend was observed for the grapes harvested from the 2001-2002 vintage, compared with the 2002-2003 vintage, although the differences were marginal.

Slightly higher populations of bacteria were found in the 2001-2002 grapes, and there was a greater incidence of C. flaccumfaciens on these grapes.

Most investigations of grapes were conducted throughout the 2002-2003 vintage, which was uncharacteristically very hot and dry. In fact, the regions from where the grapes were sampled, were officially declared as drought affected during 2002-2003.

Temperatures as high as 45qC were frequently experienced in the vineyards during the

December-February sampling period. Some general climate data for the regions where the grapes were cultivated are given in Fig. 3.6, but they do not necessarily reflect the

G G 109 unpredictable daily fluctuations in conditions that occurred from vineyard to vineyard.

Also, the reliability of the data provided by viticulturists from the various vineyards is uncertain, and the data are not complete. Thus, from a microbiological perspective, only broad conclusions can be drawn about climate influences. The hot, dry conditions of the

2002-2003 season, probably resulted in grapes with very low populations of bacteria

(102-103 CFU/g), and the general absence of lactic acid bacteria (e.g. Lactobacillus,

Pediococcus, Leuconostoc, Oenococcus)and acetic acid bacteria (e.g. Gluconobacter,

Acetobacter) that are reported in the literature to be found on grapes (Lafon-Lafourcade and Joyeux 1981; Joyeux et al. 1984; Wibowo et al. 1985; Drysdale and Fleet 1988;

Lonvaud-Funel 1999). To gain more understanding of the impact of climatic factors on the bacterial ecology of grapes, it will be necessary to conduct follow up investigations of the vineyards examined in this study, during higher rainfall vintages. The very low bacterial populations found on the grapes also prevented any sound conclusions to be drawn about the evolution of bacterial species on grapes throughout cultivation and phenological development on the vine.

With the exception of vineyard B, all vineyards followed a stringent regime of pesticide applications to control a diversity of pests. The program of pesticide application for each vineyard is given in Appendix 3. Appendix 4 provides more details about the individual pesticides applied, and the pests to be controlled. In practice, pesticides are usually applied as a mixture of several different pesticide preparations (Appendix 3).

Although there are numerous suggestions in the literature that residues of pesticides can affect the survival and growth of microflora, the incidence of wine yeasts on grapes

(van der Westhuizen et al. 2000a; 2000b) and functioning of malolactic bacteria (Vidal et al. 2001), systematic studies of these influences at the stage of grape cultivation have

G G 110 not been reported. Overall, the data of this study do not suggest any obvious impact of pesticide application on the bacterial populations or bacterial species of grapes except the overwhelming presence of B. thuringiensis, which originated from its regular use as a biological pesticide. Grapes from the organic vineyard (vineyard B), generally exhibited lower bacterial populations than those from vineyards treated with pesticides, but this correlation needs substantiation by further investigation. Ng et al. (2005) have suggested that pesticide applications can, in fact, increase the microbial load of agricultural products because bacteria may grow in some pesticide preparations.

Interestingly, B. thuringiensis was recovered from grapes samples from vineyard B, despite no application of pesticides for many years. This observation suggests a certain ubiquity of B. thuringiensis in vineyard ecosystems, possibly originating from its extensive and sustained use in viticultural practices (Bae et al. 2004).

Damaged grape berries generally exhibited higher populations and a greater diversity of bacterial species than undamaged grape berries, although the differences were not as significant as suggested by the literature (Lafon-Lafourcade and Ribéreau-Gayon 1984; du Toit and Pretorius 2000; du Toit and Lambrechts 2002). Recognized wine bacteria (G. oxydans, Acetobacter spp., Lb. kunkeei) were isolated from samples of damaged grapes, but not undamaged grapes. These conclusions were more evident for grapes sampled during the higher rainfall vintage (2001-2002) than the drier 2002-2003 vintage. Grapes exhibit damage from a variety of sources such as sun and wind damage, insect and bird damage (Buchanan and Amos 1992) and this could affect the population and diversity of associated bacteria. No attempt was made to select grapes on the basis of damage type. However, it can be stated that all damaged grapes examined had obvious physical defects to the surface layer and leaky, underlying tissue. The type of grape berry damage

G G 111 and the factors which initiate this damage could be important considerations that require more careful description and evaluation with respect to microbial ecology. Further research is needed to address these concepts as they may have an important influence on the bacterial species that occur in grape must or juice and contribute to wine fermentations.

G G 112 CHAPTER FOUR

LACTIC ACID BACTERIA ASSOCIATED WITH WINE GRAPES

4.1 INTRODUCTION

Lactic acid bacteria have a significant role in the production of wines (Lonvaud-Funel

1999; Fleet 2001, 2003a). They are responsible for conducting the malolactic fermentation (MLF) which is an important secondary reaction that occurs in many wines after the alcoholic fermentation by yeasts. The MLF has important influences on wine acidity, flavour and microbiological stability (Henick-Kling 1993, 1995; Lonvaud-

Funel 1995, 1999). Oenococcus oeni, formerly Leuconostoc oenos, is the main species responsible for MLF, although other species of lactic acid bacteria may also conduct this reaction. Various species of Lactobacillus, Pediococcus and Leuconostoc can cause spoilage of wine during bulk storage in the cellar and after bottling (Sponholz 1993; du

Toit and Pretorius 2000). Growth of lactic acid bacteria in grape juice can have inhibitory effects on yeasts and cause stuck incomplete, and sluggish alcoholic fermentations (Bisson 1999; Fleet 2003a). Despite their importance, few studies have specifically investigated the origin or source of lactic acid bacteria in the winemaking process.

Freshly extracted grape juice or must, produced under commercial conditions, generally contains various species of Lactobacillus, Pediococcus, and Leuconostoc at populations of 102-103 CFU/mL (Costello et al. 1983; Fleet et al. 1984; Pardo and ZuĔiga 1992;

Fleet 1993; Fugelsang 1997). It is considered that these bacteria originate from the

G G 113 surface of the grapes or as contaminants of winery equipment that is used to process the juice or must. Surprisingly, very few studies have focussed on the bacteria associated with grapes. Lb. plantarum, Lb. casei, Lb. brevis, Lb. hilgardii, Lb. curvatus, Lb. buchneri, Leuc. dextranicum and Leuc. mesenteroides were inconsistently isolated from several grape varieties harvested from vineyards in Spain (Sieiro et al. 1990; Suárez et al. 1994), France (Lafon-Lafourcade et al. 1983) and Germany (Weiller and Radler

1970). I have not been able to find any literature that conclusively associates the principal malolactic bacterium, O. oeni with the surfaces of either immature, mature or damaged grape berries (Radler, 1958). Nevertheless, this species is commonly isolated from wines (Chalfan et al. 1977; Davis et al. 1985, 1986; Wibowo et al. 1985; Edwards et al. 1991; Lonvaud-Funel et al. 1991).

As a part of a broad study on the yeasts and bacteria associated with wine grapes cultivated in Australia the presence of lactic acid bacteria was rarely detected by plate culture methods or by culture independent methods using PCR-denaturing gradient gel electrophoresis (DGGE) (Chapter 3). In particular, the key malolactic bacterium, O. oeni, could not be found on grapes. Possibly, these bacteria may be present on grapes at such low populations that they are not detectable by plate culture and PCR-DGGE methods. Culture enrichment methods, therefore, may be required for their isolation. In this chapter, the application of enrichment culture to investigate the presence of lactic acid bacteria on wine grapes harvested from several Australian vineyards is reported.

PCR-DGGE was also used to examine the bacterial species that developed during enrichment culture of the grapes. Both healthy, undamaged and physically damaged grape berries were analysed in this study.

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4.2 MATERIALS AND METHODS

4.2.1 Grape samples

Grape samples, collected during January - March 2004, were obtained from six commercial vineyards located in the upper and lower Hunter Valley, Mudgee and

Griffith regions of New South Wales, Australia. The northern region (upper and lower

Hunter Valley), the northwestern region (Mudgee) and the southwestern region

(Griffith) are geographically distinct from each other, being 165-595 km from Sydney.

Grapes of optimal maturity were hand-harvested and included five varieties of red grapes (Cabernet Sauvignon, Merlot, Pinot Noir, Shiraz, Tyrian) and three varieties of white grapes (Chardonnay, Sauvignon Blanc, Semillon). Among these varieties,

Cabernet Sauvignon and Shiraz from one vineyard at the upper Hunter Valley, and

Cabernet Sauvignon, Shiraz and Tyrian from one vineyard at Griffith were collected on two occasions: at 30 days prior to harvest; and at harvest time (full maturity). Each sample (about 2 kg) consisted of bunches of grapes that were randomly and aseptically removed from at least five different vines within the vineyard. Separate samples of undamaged and damaged grape bunches were taken for each variety at each vineyard.

Damaged grapes were identified on the basis of their physical appearance (broken skin, shrivelled, discoloured). Samples were stored at 4qC and analysed within 24 h of harvest from the vine.

4.2.2 Sample preparation

The overall strategy for this study is outlined in Fig. 4.1. Individual berries were randomly and aseptically removed from the bunches to give a composite sample of about 500 g. They were placed in sterile Stomacher bags and homogenized for 2 min

G 115

Grapes

Extracted grape juice

Enrichment cultures (Autoenrichment, MRS, MRS + EtOH, MRST[pH 5.5], MRST [pH 3.5])

Isolation of lactic acid bacteria by plate culture

DNA Extraction

PCR-DGGE

Amplification of DGGE band excision 16S rDNA

Sequencing of 16S rDNA

Taxonomic identification

Fig. 4.1 Experimental protocol for the analysis of lactic acid bacteria on grapes

(StomacherR Lab system, Norfolk, UK). Homogenate (10 mL) was used as inoculum for enrichment cultures.

Enrichment cultures were conducted in the following media: (i) de Man, Rogosa and

Sharpe (MRS) broth, (ii) MRS broth containing 5% (v/v) ethanol (Sigma, Chemical Co.,

St. Louis, MO.), (iii) MRST 5.5 broth which consisted of MRS broth supplemented with

15% (v/v) tomato juice (Berri, Australia), adjusted to pH 5.5, and (iv) MRST 3.5 broth adjusted to pH 3.5. MRS broth was obtained from Oxoid, Melbourne, Australia. All G 116 these media were supplemented with 100 mg/L of cycloheximide (AppliChem GmbH,

Darmstadt, Germany) to prevent the growth of yeasts and other fungi. Media were dispensed as 90 mL volumes in screw capped containers (120 mL size, Sarstedt

Australia Pty. Ltd., Technology Park, SA, Australia). While MRS broth is widely used to culture and isolate lactic acid bacteria, its performance can be improved by fortification with tomato juice (Pan et al. 1982). The conditions of 5% ethanol and pH

3.5 were included to bias the selectivity to wine conditions. An additional enrichment culture, termed autoenrichment, was conducted and consisted of a sample (100 mL) of the grape homogenate (with 100 mg/L of cycloheximide) that was incubated at 25qC.

All enrichment cultures were incubated at 25qC for 10 days. Cultures were sampled for isolation of bacteria at 0 day (no enrichment), 5 days and 10 days.

4.2.3 Isolation of lactic acid bacteria

Samples of enrichment and autoenrichment cultures were diluted as required in sterile

0.1% bacteriological peptone (Oxoid) and aliquots (0.1 mL), were spread inoculated in duplicate over the surface of plates of MRS agar or MRST agar (pH 5.5), both supplemented with cycloheximide to a final concentration of 100 mg/L. After incubation at 25qC for 3-7 days, presumptive lactic acid bacteria were selected and restreaked on the corresponding agar medium (without cycloheximide) to obtain pure cultures. Isolates were maintained as liquid cultures in MRS broth with 30% glycerol at

-80qC. Generally, the primary isolation plates from enrichment cultures gave colonies that were homogeneous in their morphology and appearance. Consequently, only two to three representative colonies were detected from enrichment samples for detailed identification.

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4.2.4 Identification of lactic acid bacteria

Isolates were presumptively classified as lactic acid bacteria if they were rods or cocci that gave Gram positive and catalase negative reactions. They were then identified by sequence analysis of segments of the 16S ribosomal DNA. Essentially, pure cultures were grown and the DNA was extracted as described later. The extracted DNA was amplified by PCR using the universal primers flanking the region between 968 and

1401 (E. coli numbering) as described by Bae et al. (2004). The amplicons were confirmed by agarose gel electrophoresis and then used directly for sequencing with

ABI PRISM£ BigDye• Terminators v3.1 Cycle Sequencing kit (Applied Biosystems,

Foster City, CA, USA). The products were sequenced at the Automated DNA analysis facility, School of Biotechnology and Biomolecular Sciences, University of New South

Wales. The sequences were aligned to determine the closest known relatives of 16S rDNA gene fragments available from BLAST in the GENBANK database (Karlin and

Altschul 1990).

Some isolates were subject to further phenotypical characterization, as an additional approach to confirm their identification. These isolates, as well as reference strains of lactic acid bacteria, were grown on MRS agar plates until prominent colonies were observed. Biomass from single colonies was mixed in sterile saline to give a suspension of 107-108 CFU/mL. This suspension was used to inoculate media for the following reactions: gas formation from glucose; production of from L-; mannitol formation from fructose (Pilone et al. 1991); growth at 10qC, 15qC, 37qC and

45qC in MRS broth; growth in MRS broth at pH 4.8 or 9.6; and growth in MRS broth containing 3.0 or 6.5% NaCl (Kandler and Weiss 1986; Schleifer 1986). Reference strains used for these tests were Lb. brevis UNSW 055100, Lb. casei subsp. rhamnosus

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UNSW 060400, Lb. plantarum UNSW 084800, Leuc. mesenteroides UNSW 060700, and Ped. pentosaceus UNSW 047200. They were obtained from the culture collection,

School of Biotechnology and Biomolecular Sciences, University of New South Wales.

4.2.5 DNA extraction

DNA was extracted by procedures described previously (Lopez et al. 2003). For isolation of total DNA from either pure cultures or samples of enrichment cultures, the cell pellet from a 2 mL culture sample (centrifuged at 10,000ug, 10 min) was resuspended in 200 ȝL of breaking buffer (2% Triton X-100, 1% SDS, 100 mM NaCl,

10 mM Tris-Cl [pH 8.0], 1 mM EDTA [pH 8.0]) and 20 ȝL of lysozyme (10 mg/mL,

Sigma) and incubated for 15 min at 37qC. The mixture was transferred to a micro tube containing 0.3 g of 0.1 mm diameter zirconia/silica beads (BioSpec Products Inc.,

Bartlesville, OK). The cells were homogenized twice for 40 s at a speed setting of 46 in a bead beater (Mini-BeadBeater-1™, BioSpec Product Inc.) in the presence of 200 ȝL of phenol-chloroform-isoamylalcohol (25:24:1). Two hundred microliters of TE buffer

(10 mM Tris-Cl, 1 mM EDTA, pH 8.0) were then added, and the bead cell mixture was centrifuged at 10,000ug for 10 min at 4qC. The aqueous phase was transferred to another microcentrifuge tube, and the DNA was precipitated by the addition of 0.6 volume of isopropanol, and recovered by centrifugation at 10,000ug for 10 min at 4qC.

The DNA pellet was washed with 70% ethanol, dried and resuspended in 50 ȝL of sterile TE buffer. The crude DNA extracts of enrichment cultures were further purified by the Wizard DNA purification kit (Promega Corp., Madison, WI, USA) according to the protocol supplied by the manufacturer.

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4.2.6 Analysis of enrichment cultures by PCR-denaturing gradient gel

electrophoresis (DGGE)

Purified DNA was amplified with primers 341f-gc/518r (Muyzer et al. 1993) and 968f- gc/1378r (Heuer et al. 1999) spanning the V3 region and V6-V9 region of the 16S ribosomal DNA, respectively (Table 4.1). The 40 nucleotide GC rich sequence at the 5’ end of the forward primers (341f and 968f) was attached to improve the detection of sequence variations of amplified DNA fragments by subsequent DGGE (Muyzer et al.

1993). These primers were obtained from SigmaGenosys, Australia. The PCR reaction mixtures contained 10 mM Tris-HCl [pH 8.3], 50 mM KCl, 0.2 ȝM of each primer, 200

ȝM of each dNTP (Roche Diagnostics, Indianapolis, IN, USA), 1.5 mM MgCl2, 1.25 U of Gold Taq DNA polymerase (AmpliTaq•, Roche Molecular Systems, Branchburg, NJ,

USA) and 10 ng of purified template DNA in 50 ȝL final volume. The concentration of

MgCl2 was increased to 3.75mM for PCR with the 968f-gc/1378r. PCR amplifications were conducted in a 9600 Thermal Cycler (Applied Biosystems). The PCR amplification products were verified by 2% agarose gel electrophoresis, and then resolved by DGGE using a DCode• Universal Mutation Detection System (Bio-Rad,

Hercules, CA, USA) to determine the species composition of enrichment cultures.

Essentially, DGGE was performed as described by Muyzer et al. (1993). Eight percent polyacrylamide gels (acrylamide:N,N’-methylene bisacrylamide ratio, 19:1, Bio-Rad) with 30-50% linear denaturing gradient (100% denaturant corresponds to 7 M urea and

40% formamide) were used for analysing amplicons of 341f-gc/518r. Electrophoresis was performed for 30 min at 20 V and for 6 h at 120 V with a constant temperature of

60qC. Six percent polyacrylamide gels with 45-65% linear denaturing gradient were used for analysing amplicons produced with primers of 968f-gc/1378r, and electrophoresis was run for 30 min at 20 V, then for 14 h at 100 V. Subsequently, gels

G Table 4.1 Primers and PCR amplification conditions used in this study Primer Sequence (5'-3') PCR amplification condition Initial Main cycle Final denaturation extension ab 341 f -gc CCTACGGGAGGCAGCAG o o o o Eubacteria V3 (341-357) 95 C, 7 min 94 C, 1 min 94 C, 1min 72 C, 10min 65~55o C,d 1min 20 cycles 55o C, 1min 10 cycles 518r ATTACCGCGGCTGCTGG o o Universal V3 (518-534) 72 C, 1min 72 C, 1min

968f-gc AACGCGAAGAACCTTAC o o o o Eubacteria V6 (968-984) 95 C, 7 min 94 C, 1 min 94 C, 1min 72 C, 10min 60~55o C,d 1min 10 cycles 55o C, 1min 20 cycles 1378r CGGTGTGTACAAGGCCC- 72o C, 2min 72o C, 2min Eubacteria V9 (1378-1401) GGGAACG GC clampc cgcccgccgcgccccgcgcccgtcccgccgccccgcccg aBase numbered relative to E. coli 16S rRNA sequence bf, forward primer; r, reverse primer cGC clamp added to the 5' end of the forward primers d1o C touch down every second cycle 120 G 121 were stained with SYBR“ Green I (Amresco“, Ohio, USA) for 20 min in 1u TAE buffer (40 mM Tris-acetate, 1 mM EDTA, pH 8.0) and photographed under UV transillumination with a Polaroid DS-34 camera.

DNA bands in DGGE gels were excised and incubated overnight in 50 ȝL of sterile

Milli-Q water at 4qC. Eluted DNA was reamplified with corresponding primers and the

PCR products were re-evaluated by DGGE. Only products that migrated as a single band and at the same distance with respect to the original sample were then excised and amplified with corresponding primers without a GC clamp. Amplified products were purified with QIAquick PCR purification kit (Qiagen Pty Ltd., Clifton Hill, Australia), and subjected to DNA sequence analysis.

4.2.7 Interactive growth of Oenococcus oeni, Lactobacillus kunkeei and

Lactobacillus lindneri.

To investigate interactive growth between O. oeni and Lb. kunkeei, and O. oeni and Lb. lindneri, mixed cultures were conducted in MRST broth at pH 3.5 and pH 5.5. Lb. kunkeei and Lb. lindneri were isolated from samples of enrichment and autoenrichment cultures. Two strains of O. oeni were isolated by plating samples from commercial starter culture products onto MRST agar. O. oeni 1 and O. oeni 2 were isolated from

Viniflora CH35• and Viniflora Oenos• (CHR. Hansen Pty. Ltd., Australia) respectively. O. oeni, grown in MRST broth at 25qC for 5 days, and Lb. kunkeei or Lb. lindneri, grown in MRST broth at 25qC for 24 h, were inoculated as single or mixed cultures into 200 mL of MRST broth in a 250 mL Duran“ Schott bottle (Schott UK Ltd.,

UK). The cultures were incubated at 25qC for 10 days. Samples were removed every 12

G G 122 h or 24 h to determine the population of these microorganisms by spreading plating onto

MRST agar. Isolates of O. oeni, Lb. kunkeei and Lb. lindneri were also characterised for their interactions using the spot on lawn assay and the deferred antagonism assay as described previously (Bae et al. 2004).

4.3 RESULTS

4.3.1 Isolation of lactic acid bacteria from enrichment cultures

To increase the prospects of isolating lactic acid bacteria, the experimental plan included analysis of grapes representing eight different cultivars, grapes harvested from six vineyards located in four different wine producing regions, and samples of undamaged and damaged grape berries. Overall, 43 batches of grapes (21 undamaged,

22 damaged) were examined. Samples from each batch were analysed by five different enrichment procedures. Isolations were prepared from each enrichment culture after 5 days of incubation (when microbial growth first became evident) and, again, after 10 days of incubation. Tables 4.2-4.5 show the species of lactic acid bacteria isolated from grapes according to this plan. The populations of the species in the enrichment cultures are given in the Tables. With a few exceptions, they were in the range 107-109 CFU/mL.

All grape samples were analysed by direct plate culture before enrichment (0 days), but only a few samples of damaged grapes of Chardonnay (vineyard D), Shiraz and

Chardonnay (vineyard E) and Sauvignon Blanc (vineyard F) gave isolation of lactic acid bacteria (Lb. lindneri, Lb. kunkeei) at 102-104 CFU/mL by this method.

Lactic acid bacteria were isolated from only 9 of the 21 batches of undamaged grapes and 16 of 22 batches of damaged grapes (Tables 4.2-4.5). Thus, there were numerous

G G 123 batches from which no lactic acid bacteria could be cultured. In many cases, bacteria other than lactic acid bacteria were found in enrichment cultures. For example, Cabernet

Sauvignon and Tyrian grapes from vineyard F of the Griffith region (Table 4.2) gave no lactic acid bacteria in any of the enrichment cultures, but Sporolactobacillus inulinus

(Gram positive rod, spore former, catalase negative and identified by sequencing) was isolated in MRS, MRS+EtOH and MRST pH 5.5 enrichment cultures at populations of

105-107 CFU/mL. This species was also recovered from some samples of Shiraz and

Semillon grapes in other vineyards. No lactic acid bacteria were isolated from undamaged Semillon grapes from vineyard A of the Hunter Valley region (Table 4.5) but all enrichment cultures, except MRST+EtOH, showed the presence of Asaia siamensis (Gram negative rod, catalase positive and identified by sequencing). This species was also found on some Shiraz grapes of vineyards B and F. Species of Bacillus,

Staphylococcus, and Gluconobacter were also detected infrequently. Non-lactic acid bacteria were mostly detected in enrichment cultures where lactic acid bacteria were not present, but there were some enrichment cultures, that contained both non-lactic acid bacteria and lactic acid bacteria. Overall, very few batches (2 out of 21 undamaged grapes and 1 out of 22 damaged grapes) showed no bacterial presence in enrichment cultures. Detailed analyses and identification of the non-lactic acid bacteria were beyond the focus and scope of this Chapter. The data observed are presented in Tables

5.1-5.5 of Appendix 5 and form the basis for future investigation.

Table 4.6 lists the different species of lactic acid bacteria that were found and their frequency of isolation from the different batches of grapes. The frequency of isolation was higher from white grape varieties (25/19, isolation/batches) than from red grape varieties (14/24), and from damaged grape samples (27/22) than from undamaged grape

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Table 4.2 Isolation of lactic acid bacteria by enrichment culture from Cabernet Sauvignon, Merlot, Pinot Noir and Tyrian grapes

Enrichment condition Grape Physical Vineyarda Autoenrichment MRS MRS + EtOH MRST (pH 5.5) MRST (pH 3.5) variety statusb 5 d 10 d 5 d 10 d 5 d 10 d 5 d 10 d 5 d 10 d Cabernet A UD ------Sauvignon D Lb. mali Lb. mali Lb. Lb. Lb. mali Lb. Lb. mali Lb. mali Lb. Lb. mali (6.4u107)c (2.4u108) plantarum plantarum (4.0u108) plantarum (1.0u108) (2.3u108) plantarum (2.0u106) Lb. Lb. (5.0u108) (4.1u108) Lb. (2.8u108) Lb. Lb. (2.5u108) Lb. plantarum plantarum plantarum plantarum plantarum plantarum (2.8u106) (1.4u107) (9.0u108) (6.7u108) (2.8u108) (2.4u107) D UD ------D - - Lb. lindneri Lb. lindneri ------(2.7u109) (4.3u108) F UD ------D ------Merlot D UD - - Lb. lindneri Lb. lindneri - Lb. lindneri Lb. lindneri Lb. lindneri - - (2.9u109) (1.8u108) (4.0u105) (7.5u108) (1.2u107) D - - E. durans E. durans E. durans E. durans - - - - (1.3u108) (1.0u108) (1.6u106) (6.5u108) Pinot D UD ------E. durans E. durans - - Noir (1.0u105) (3.6u107) D ------Tyrian F UD ------D ------a Vineyard location; A, Hunter Valley; D, Mudgee; F, Griffith b UD, undamaged grape berries; D, damaged grape berries c population (CFU/mL) in enrichment culture

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Table 4.3 Isolation of lactic acid bacteria by enrichment culture from Shiraz grapes

Enrichment condition Physical Vineyarda Autoenrichment MRS MRS + EtOH MRST (pH 5.5) MRST (pH 3.5) statusb 5 d 10 d 5 d 10 d 5 d 10 d 5 d 10 d 5 d 10 d A UD - - Lc. lactis Lc. lactis - - Lc. lactis Lc. lactis - - (2.1u109)c (2.1u108) (1.9u109) (5.9u107) Lb. lindneri Lb. lindneri (3.0u106) (3.0u107) D - - - - Lc. lactis Lc. lactis - - - - (6.8u108) (1.3u104) B UD ------D - - Lb. lindneri Lb. lindneri Lb. lindneri Lb. lindneri Lb. lindneri Lb. lindneri - - (2.8u108) (5.6u107) (9.0u108) (7.0u108) (2.8u109) (3.4u108) C UD - - E. avium E. avium - E. avium E. avium E. avium - - (1.7u107) (6.8u108) (3.7u105) (2.5u103) (7.3u106) D ------D UD ------D ------E UD ------D - - Lb. lindneri Lb. lindneri Lb. lindneri Lb. lindneri Lb. lindneri Lb. lindneri Lb. lindneri Lb. lindneri (4.0u108) (8.4u108) (3.0u108) (6.8u108) (1.0u108) (1.8u109) (4.3u102) (2.1u104) E. durans E. durans (7.0u105) (1.3u106) F UD ------D - - Lc. lactis Lc. lactis Lc. lactis Lc. lactis - - - - (1.7u109) (3.6u106) (4.3u103) (8.6u108) a Vineyard location; A, B, C, Hunter Valley; D, E, Mudgee; F, Griffith b UD, undamaged grape berries; D, damaged grape berries c population (CFU/mL) in enrichment culture

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Table 4.4 Isolation of lactic acid bacteria by enrichment culture from Chardonnay grapes

Enrichment condition Physical Vineyarda Autoenrichment MRS MRS + EtOH MRST (pH 5.5) MRST (pH 3.5) statusb 5 d 10 d 5 d 10 d 5 d 10 d 5 d 10 d 5 d 10 d A UD - - E. avium E. avium E. avium E. avium E. avium E. avium - - (1.0u107) (8.8u108) (1.0u107) (4.5u108) (1.0u106) (6.2u108) D ------C UD Lb. kunkeei Lb. kunkeei E. faecium E. faecium - - Lb. kunkeei Lb. kunkeei Lb. kunkeei Lb. kunkeei (1.0u107)c (1.6u104) (4.2u107) (1.0u108) (2.7u106) (4.0u106) (1.0u105) (2.7u106) D - - Lc. lactis L.lactis Lb. kunkeei Lb. kunkeei Lb. lindneri Lb. lindneri - - (1.0u107) (3.0u108) (3.2u106) (2.9u106) (2.7u107) (7.0u107) D UD - - E. durans E. durans E. durans E. durans E. durans E. durans - - (1.0u107) (1.7u106) (8.1u102) (5.4u107) (5.0u107) (2.7u107) D Lb. lindneri Lb. lindneri Lb. lindneri Lb. lindneri Lb. lindneri Lb. lindneri Lb. lindneri Lb. lindneri Lb. lindneri Lb. lindneri (3.8u108) (3.2u108) (2.2u109) (5.2u108) (1.5u109) (7.0u108) (2.8u109) (1.3u109) (3.4u106) (1.1u109) E UD ------D Lb. kunkeei Lb. kunkeei Lb. lindneri Lb. lindneri Lb. lindneri Lb. lindneri Lb. lindneri Lb. lindneri Lb. lindneri Lb. lindneri (1.0u107) (1.6u104) (1.1u107) (1.6u104) (7.8u108) (4.5u107) (1.5u109) (9.5u107) (7.8u107) (3.2u108) F UD - - W. W. E. faecium E. faecium - - - - paramesenter paramesenter (6.8u108) (5.5u106) oides oides W. W. (1.3u108) (1.0u104) paramesenter paramesenter oides oides (1.3u108) (1.0u104) D - - E. durans E. durans W. W. Lactobacillus Lactobacillus - - (7.8u108) (5.2u108) paramesenter paramesenter sp. sp. oides oides (9.5u109) (6.7u109) (1.3u108) (1.0u104) a Vineyard location; A, C, Hunter Valley; D, E, Mudgee; F, Griffith b UD, undamaged grape berries; D, damaged grape berries c population (CFU/mL) in enrichment culture

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Table 4.5 Isolation of lactic acid bacteria by enrichment culture from Semillon and Sauvignon Blanc grapes

Enrichment condition Grape Physical Vineyarda Autoenrichment MRS MRS + EtOH MRST (pH 5.5) MRST (pH 3.5) variety statusb 5 d 10 d 5 d 10 d 5 d 10 d 5 d 10 d 5 d 10 d Semillon A UD ------D - - E. her- E. her- E. her- E. her- - - - - manniensis manniensis manniensis manniensis (1.0u107) c (1.8u107) (1.0u107) (7.2u107) B UD ------D - - Lb. lindneri Lb. lindneri Lb. lindneri Lb. lindneri Lb. lindneri Lb. lindneri - - (3.0u108) (1.0u108) (3.0u108) (9.0u107) (1.0u108) (1.7u107) F UD - - - -W. para-W. para- - - - - mesenteroides mesenteroides (8.0u107) (8.0u104) D - - W. para- Lb. kefiri W. para- E. faecium W. para- E. faecium - - mesenteroides (2.0u107) mesenteroides (4.0u104) mesenteroides (5.3u104) (1.6u108) (4.0u107) W. para- (4.4u107) W. para- mesenteroides mesenteroides (2.0u104) (4.0u103) Sauvignon D UD ------Blanc D Lc. lactis - E. durans E. durans E. durans E. durans - - - - (1.0u107) (1.7u109) (4.4u108) (9.6u108) (2.4u108) F D Lb. Lb. Lb. kunkeei Lb. kunkeei Lb. kunkeei Lb. kunkeei Lb. kunkeei Lb. kunkeei Lb. Lb. kunkeei kunkeei (1.4u108) (8.0u107) (9.6u106) (4.0u107) (4.4u107) (9.0u107) kunkeei kunkeei (3.5u108) (1.2u109) Lb. lindneri Lb. lindneri Lb. lindneri Lb. lindneri Lb. lindneri Lb. lindneri (4.2u107) (3.1u104) (1.4u108) (1.2u108) (1.6u107) (1.2u108) (2.4u107) (2.2u108) Lb. Lb. lindneri lindneri (4.1u107) (3.6u104) a Vineyard location; A, B, Hunter Valley; D, Mudgee; F, Griffith b UD, undamaged grape berries; D, damaged grape berries c population (CFU/mL) in enrichment culture

G G 128 samples (12/21). The isolates consisted of six species of Lactobacillus, four species of

Enterococcus, and one species from each of Lactococcus and Weissella. Species of

Leuconostoc, Pediococcus and Oenococcus were not found. The most frequently isolated species were Lb. lindneri, E. durans, Lc. lactis and Lb. kunkeei. The data are too limited to show any definitive relationship between species occurrence, grape variety and vineyard from where the grapes were collected.

Several batches of red grapes were collected from vineyards A and F, 30 days before their scheduled harvest time. For vineyard A, associated lactic acid bacteria and grapes were E. hermanniensis (Shiraz), and E. avium, E. hermanniensis, Lb. lindneri (Cabernet

Sauvignon), and for vineyard F, the data were E. faecium, W. paramesenteroides

(Shiraz), E. faecium (Cabernet Sauvignon) and W. paramesenteroides (Tyrian).

Table 4.6 Frequency of isolation of lactic acid bacteria from undamaged and damaged wine grapes

Undamaged grapes Damaged grapes Species Total Ra (12) b W (9) R (12) W (10) Lb. lindneri 2 0 3 5 10 E. durans 1 1 2 2 6 Lc. lactis 1 0 2 2 5 Lb. kunkeei 0 1 0 3 4 W. paramesenteroides 0 2 0 2 4 E. faecium 0 2 0 1 3 E. avium 1 1 0 0 2 Lb. kefiri 0 0 0 1 1 Lb. mali 0 0 1 0 1 Lb. plantarum 0 0 1 0 1 Lactobacillus sp. 0 0 0 1 1 E. hermanniensis 0 0 0 1 1 Total 5 7 9 18 39 a R, red grape variety; W, white grape variety, b number of batches

The reasons for using an array of enrichment conditions were described previously (see also Pan et al. 1982). It is evident from Tables 4.2-4.5 that, for some grape batches, the enrichment medium could determine whether or not any lactic acid bacteria were

G G 129 isolated at all, and the species diversity found. Autoenrichment, and enrichment in

MRST adjusted to pH 3.5 were the conditions that gave the least isolations of lactic acid bacteria (Table 4.7). Similar, overall isolation frequencies were found for enrichment in

MRS, MRS+EtOH and MRST pH 5.5, but there were examples where some species were isolated under one condition but not the other. For damaged Chardonnay grapes from vineyard C, Lc. lactis was isolated by enrichment in MRS, Lb. kunkeei was obtained in MRS+EtOH, and Lb. lindneri was found in MRST pH5.5 (Table 4.4).

Damaged Chardonnay grapes from vineyard F gave E. durans in MRS, W. paramesenteroides in MRS+EtOH, and Lactobacillus spp. in MRST pH 5.5. While most samples gave only one species after enrichment, there were some samples where more than one species was found (e.g. Lb. mali and Lb. plantarum in damaged Cabernet

Sauvignon from vineyard A, Table 4.2; Lb. kunkeei and Lb. lindneri in damaged

Sauvignon Blanc from vineyard F, Table 4.5).

Table 4.7 Frequency of isolation of lactic acid bacteria from wine grapes according to enrichment medium

Species isolated Enrichment condition Autoenrichment MRS MRS + EtOH MRST, MRST Total (pH 5.5) (pH 3.5) Lb. lindneri 1 9 7 8 4 29 E. durans 0 5 4 2 0 11 Lb. kunkeei 3 1 2 2 2 10 Lc. lactis 1 3 2 1 0 7 W. paramesenteroides 0 2 4 1 0 7 E. avium 0 2 2 2 0 6 Lb. plantarum 1 1 1 1 1 5 Lb. mali 1 0 1 1 1 4 E. faecium 0 1 2 1 0 4 E. hermanniensis 0 1 1 0 0 3 Lactobacillus sp. 0 0 0 1 0 1 Lb. kefiri 0 1 0 0 0 1 Total 7 26 26 20 8 87

4.3.2 Identification of isolates

A total of 160 strains were isolated from the enrichment cultures. All isolates were

G G 130 considered to be lactic acid bacteria based on their positive Gram reactions, non-motility, absence of catalase activity and spore formation, and rod or coccal shape. All 160 isolates were identified by sequencing of partial 16S ribosomal DNA and then divided into nine major groups based on sequence analysis. Representative isolates from each group were examined for several key phenotypical tests listed in Table 4.8.

Group I included 13 strains showing ovoid cells in short chains that did not produce gas from glucose and hydrolyse L-arginine. They did not grow at 45qC, which is different from the standard description, and gave weak growth at pH 4.8 after incubation for 7 days. The identity of all 13 strains was determined as E. avium (AY442814) with 96-

99% DNA sequence similarity. Groups II, III and IV were homofermentative cocci in short chains and hydrolysed L-arginine to ammonia. In comparison with other groups, isolates of group II grew well over a range of different conditions. All fourteen strains were identified as E. faecium (AJ420800) with 97-99% homology. No growth at pH 4.8 and relatively weak growth at 45qC were detected for isolates of group III. All 14 isolates of group III were shown to be E. durans (AJ420801) with 97-99% homology.

Group IV included six strains that did not grow at 45qC, which is unlike the other enterococci, and showed 98-99% DNA sequence similarity to E. hermanniensis.

Group V included seven strains with similarity to enterococci, but they were distinguished by their inability to grow at 6.5% NaCl and pH 9.6. They were shown to be Lc. lactis (AF493058) with 99% DNA sequence homology.

Group VI isolates were short rods, produced gas from glucose and gave no formation of ammonia from L-arginine, and mannitol from fructose. They did not grow at 10qC or

45qC. A total of 11 strains of group V were identified as W. paramesenteroides

(S67831) with 97 - 99% sequence similarity.

G Table 4.8 Characteristics of lactic acid bacteria isolated from wine grapes

Characteristic Group I Group IIGroup IIIGroup IV Group V Group VI Group VII Group VIII Group IX

No. isolates 13 14 14 6 7 11 20 58 7 No. isolates tested 2 2 5 1 1 2 6 9 2 Identity by sequencing E. avium E. faecium E. durans E. hermanniensis Lc. lactis W. paramesenteroides Lb. kunkeei Lb. lindneri Lb. plantarum Accession No. AY442814 AJ420800 A420801 AY396048 AF493058 S67831 Y11374 X95421/D37784 AL935260 Homology 96-99% 97-99% 97-99% 98-99% 99% 97-99% 98-99% 97-98% 98-99%

Cellular morphology cocci cocci cocci cocci cocci short rod rod rod rod Gram reaction + (+)a1 + (+)1 + (+)1 + (+)2 + (+)1 + (+)3 + (+)4 + (+)5 + (+)6 Catalase test - (-) - (-) - (-) - (-) - (-) - (-) +/- (+/-) +/- (-) - (-) Gas from glucose - (-) - (-) - (-) - (-) - (-) + (+) + (+) + (+) - (-) of L-Arginine - (-) +/- (+) + (+) + (+) + (+) - (-) - (-) - (-) - (-) Mannitol Formation - (-) - (-) - (-) - (-) - (-) - (-) + (+) + (+) - (-) Growth at temperature 10o C + (d) + (+) + (+) + (+) + (+) - (-) - (-) + (+) + (+) 15o C + (+) + (+) + (+) + (+) + (+) + (+) + (+) + (+) + (+) 37o C + (+) + (+) + (+) + (+) + (+) + (+) + (+) + (+) + (+) 45o C - (+) + (+) +/- (+) - (-) - (-) - (-) - (-) - (-) - (-) Growth in NaCl 3.0 % + (+) + (+) + (+) + (+) + (+) + (+) + (+) + (+) + (+) 6.5 % + (d) + (+) + (+) + (V) - (-) + (+) - (-) + (+) + (+) Growth at pH 4.8 +/- (ND) + (ND) - (ND) - (ND) + (+) + (+) + (+) + (+) + (+) 9.6 + (+) + (+) + (+) + (+) - (-) + (+) - (-) - (-) - (-) +, growth; -, no growth; +/-, weak growth or weak reaction; d, delayed reaction; v, variable reaction; ND, no data. a , strandard description of reference strains;1 , Schleifer 1986;2 , Koort et al. 2004;34 , Cai et al. 1998; , Edwards et al. 1998b; 5 , Back et al. 1996; 6, Kandler and Weiss 1986. 131 G 132

A large number of lactic acid bacteria (groups VII and VIII) were distinguished by their rod cell shape, presence of weak catalase activity, production of gas from glucose and mannitol from fructose, and their inability to hydrolyse L-arginine. Group VII showed different growth patterns at 10qC and 6.5% NaCl compared with active growth of group

VIII at these conditions. All isolates of group VII were assigned as Lb. kunkeei

(Y11374) with 98 - 99% sequence similarity, and isolates of group VIII were identified as Lb. lindneri (X95421 and D37784) with 97 - 98% DNA sequence homology.

Isolates in group IX differed from the other groups by their rod shape, homofermentative property, and failure to produce mannitol from fructose. They grew well under most conditions except at 45qC and pH 9.6. Sequence analysis identified this group as Lb. plantarum (AL935260) with 98 - 99% similarity.

4.3.3 Analysis of lactic acid bacteria in enrichment cultures by PCR-

DGGE

In addition to plating on agar media, PCR-DGGE was used to detect lactic acid bacteria in several of the enrichment cultures reported in Tables 4.2-4.5. Fig. 4.2 shows the PCR-

DGGE banding patterns of pure cultures of various species of lactic acid bacteria isolated from grapes by enrichment-agar plating, as well as some additional reference species. Good separation of the different species was obtained. The 16S rDNA amplicons examined in Fig. 4.2 were generated by PCR using primers 341f-gc and 518r.

Amplicons produced with primers 968f-gc and 1378r were not well isolated by DGGE, even after varying the concentration of denaturant used in preparation of the electrophoresis gel (Fig. 4.3).

G 133

12345M1 678910111213 M2

Fig. 4.2 DGGE of the 16S rDNA PCR amplicons of lactic acid bacteria. Lanes: 1, Lactobacillus plantarum UNSW 084800; 2, Oenococcus oeni 1 (Viniflora CH35) 3, Lactobacillus brevis UNSW 055100; 4, Pediococcus pentosaceus UNSW 047200; 5, Lactobacillus casei UNSW 060400; M1, mixture of species in lanes 1-5; 6, Lactobacillus kunkeei;7,Weissella paramesenteroides;8,Enterococcus avium; 9, Enterococcus facium; 10, Lactobacillus kefiri;11,Enterococcus durans; 12, Lactococcus lactis; 13, Lactobacillus lindneri; M2, mixture of species in lane 6-13.

Lk Lk, W W, Lbc Lbc, Lp, Oe Lp, Oe

Pp Lb Ll Lbc Ea Lk Oe Lp Pp W Pp Lb Ll Ll,Lb Ea Ea

35-55%* 40-60% 40-52.5% 100V 14hrs 100V 14hrs 100V 14hrs

Fig. 4.3 DGGE banding patterns of 16S rDNA PCR amplicons generated with 968f-gc and 1378r primers of lactic acid bacteria . Ea, E. avium; Lb, Lb. brevis UNSW 055100; Lbc, Lb. casei UNSW 060400; Lk, Lb. kunkeei; Ll, Lb. lindneri; Oe, O. oeni; Pp, P. pentosaceus UNSW 047200; W, W. paramesenteroides. *, Gradient conditions for concentration of denaturant, electrophoretic conditions. G 134

The enrichment cultures from six batches of grapes were analysed for lactic acid bacteria by both plate culture and PCR-DGGE. For each batch, the 10 day enrichment cultures in MRS, MRS+EtOH, MRST 5.5 and MRST 3.5 were examined.

Table 4.9 shows the species detected by each method, and Fig. 4.4 shows the corresponding PCR-DGGE data. For most cases, the same species of lactic acid bacteria were detected by both methods. However, some exceptions occurred. In some examples, species of lactic acid bacteria were recovered by plating but not by DGGE. For the

Cabernet Sauvignon grapes (vineyard A), Lb. plantarum and Lb. mali were not detected by PCR-DGGE in the MRST 5.5 enrichment but were found by plating. E. faecium was not detected by PCR-DGGE but was found by plating in MRS + EtOH and MRST 5.5 enrichment of Semillon (vineyard F). For Shiraz (vineyard E) and Sauvignon Blanc

(vineyard F) grapes, Lb. lindneri was not found by PCR-DGGE in the MRST 3.5 enrichment but was recovered by plating. In other cases, lactic acid bacteria were detected by PCR-DGGE but not by plating. For example, Lc. lactis was found in enrichment cultures of the Cabernet Sauvignon (vineyard A) by PCR-DGGE but not by plate culture. In several cases, multiple bands on DGGE gels were obtained for some species. Lb. lindneri from enrichment cultures of Shiraz grapes (vineyard E) gave three bands (Fig. 4.4b) and Lb. lindneri from all enrichment cultures of Chardonnay grapes

(vineyard D) gave 9 bands (Fig. 4.4c). W. paramesenteroides from Semillon grapes

(vineyard F) gave two bands only for MRST 5.5 enrichment culture (Fig. 4.4e).

G G 135

Table 4.9 Comparison of agar plating and PCR-DGGE methods for detection of lactic acid bacteria in enrichment cultures of wine grapes a

Enrichment-plating PCR-DGGE DGGE profile b Grape sample Species 1 c 2 3 4 1 c 2 3 4 (a) Cabernet Sauvignon, Lb. plantarum + + + + + + - + damaged, vineyard A Lb. mali - - + + - + - + Lc. lactis - - - - + + + - (b) Shiraz, Lb. lindneri + + + + + + + - damaged, vineyard E (c) Chardonnay, Lb. lindneri + + + + + + + + damaged, vineyard D (d) Chardonnay, Lb. lindneri + + + + + + + + damaged, vineyard E (e) Semillon, W. paramesenteroides - + + - + + + - damaged, vineyard F Lb. kefiri + - - - + - - - E. faecium - + + - - - - - (f) Sauvignon Blanc, Lb. kunkeei + + + + + + + + damaged, vineyard F Lb. lindneri + + + + + + + - a See Tables 4.2-45 b Profiles shown in Fig.4.4 c Enrichment cultures; 1, MRS; 2, MRS + 5% Ethanol; 3, MRST pH 5.5; 4, MRST pH 3.5

G (a)(b) (c) (d)(e) (f)

M2 M1 1 2 3 4 M2 M1 1 2 3 1 2 3 4 M2 M1 M2 M1 1 2 3 4 M2 M1 1 2 3M2 M1 1 2 3 4

Lk Wp Lp

Wp

Ll Lkf Lm Ll Lc Ll Ll Ll Ll Ll

Fig. 4.4 DGGE profiles of PCR amplicons for lactic acid bacteria found in enrichment culture of grapes. Lanes: 1, MRS enrichment; 2, MRS + EtOH enrichment; 3, MRST (pH 5.5) enrichment; 4, MRST (pH 3.5) enrichment; M1 and M2, referred in Fig. 4.2. Lp, Lb. plantarum; Lm, Lb. mali; Lc, Lc. lactis; Ll, Lb. lindneri; Wp, W. paramesenteroides; Lkf, Lb. kefiri; Lk, Lb. kunkeei. (a), (b), (c), (d), (e), (f), cultures of wine grapes, referred to in Table 4.9. 136 137

The nine DNA bands obtained after PCR-DGGE analysis of MRS enrichment culture of damaged Chardonnay grapes (vineyard D) (Fig. 4.4c, lane 1; Fig. 4.5) were excised and sequenced. All of the bands gave sequences that were homologous (92-95%) with Lb. lindneri. After excision, each DNA band was subjected to another round of PCR-DGGE analysis, and these also gave multiple DNA bands that corresponded with patterns in the first analysis (Fig. 4.5, lanes 2-4). These bands all sequenced as Lb. lindneri. However, band d, from the original analysis only give one band after the second PCR-DGGE (Fig.

4.5, lane 5). The mobility of this band was the same as the original band d and corresponded to the band mobility obtained on PCR-DGGE analysis of a pure culture of

Lb. lindneri.

M1 1 2345

a b c d

Fig. 4.5 Re-evaluation of DNA bands from PCR-DGGE analysis of damaged Chardonnay grapes, vineyard D. Bands a, b, c, d from lane 1 were excised, and re-examined by PCR-DGGE giving profiles in lanes 2, 3, 4, 5, respectively. 138

Using pure cultures of either Lb. lindneri or Lb. kunkeei, it was determined that PCR-

DGGE could detect cell populations down to 103 CFU/mL. No DNA bands were obtained when PCR-DGGE was done with the population of cells diluted to 10 2

CFU/mL (data not shown). A mixture of 105 CFU/mL of Lb. kunkeei and 105 CFU/mL of Lb. lindneri gave two DNA bands after PCR-DGGE analysis although the band for

Lb. lindneri had weaker intensity (Fig.4.6, lanes 2, 6). A mixed population of Lb. kunkeei (1065 CFU/mL) and Lb. lindneri (10 CFU/mL) gave only one band with mobility of Lb. kunkeei after PCR-DGGE analysis (Fig. 4.6, lane 3), while a mixture of

Lb. lindneri (106 CFU/mL) and Lb. kunkeei (105 CFU/mL) gave two bands, but the band of Lb. kunkeei was weak (Fig. 4.6, lane 7). Thus, the PCR-DGGE assay could not reliably detect one species in a mixture if its population was 10 fold less than the other species.

M212 34 M2 56 78 Lb.kunkeei

Lb.lindneri

Fig. 4.6 PCR-DGGE detection of Lactobacillus kunkeei and Lactobacillus lindneri in mixed cell suspensions at differnt population ratios. Lane: 1, Lb. kunkeei (105 CFU/mL); Lanes: 2-4, Lb. kunkeei (CFU/mL) : Lb. lindneri (CFU/mL) 105 :1056 ; 10 :1055 ; 10 7 :10 , respectively; Lane 5, Lb. lindneri (105 CFU/mL); Lane 6-8, Lb. kunkeei (CFU/mL) : Lb. lindneri (CFU/mL) 10567 :1055 ; 10 :10 ; 10 5 :10 , respectively. G 139

4.3.4 Interactive growth of O. oeni and lactic acid bacteria

Failure to isolate O. oeni from the grapes by enrichment culture could mean (i) that it is not present in the habitat or (ii) that it could be present but is out-competed and overgrown by other, faster growing species during enrichment. To test this possibility, the growth behaviours of two strains of O. oeni in relation to two other bacteria, Lb. lindneri and Lb. kunkeei, that were frequently found on grapes were investigated. Both strains of O. oeni gave similar responses, so data for one strain only are reported. Fig.

4.7 shows the growth responses of O. oeni, Lb. kunkeei and Lb. lindneri as single cultures and when grown in mixed cultures of (i) O. oeni with Lb. kunkeei and (ii) O. oeni with Lb. lindneri. As single cultures, O. oeni grew more slowly than Lb. kunkeei or

Lb. lindneri. O. oeni failed to grow in mixed culture with Lb. lindneri and exhibited delayed growth in mixed culture with Lb. kunkeei of pH 5.5 (Fig. 4.7a). Interestingly, viability of the inoculated cells of O. oeni rapidly decreased to non-detectable levels on mixture with either Lb. kunkeei or Lb. lindneri. Such rapid loss of viability of O. oeni was not observed for the mixture at pH 3.5 (Fig. 4.7b). In fact, there was slight growth of O. oeni during the first 2 days, before it died off. However, it did not recover in the mixture with Lb. lindneri, but eventually re-established growth in the presence of Lb. kunkeei. The growth of either Lb. lindneri or Lb. kunkeei was not affected (cf. control) by co-culture with O. oeni (Fig. 4.7c, 4.7d).

The interaction of O. oeni with two species of Lactobacillus was also determined by the spot on lawn assay and deferred antagonism plate assay (see Chapter 5, Bae et al. 2004).

The two strains of O. oeni were not inhibited by Lb. kunkeei, but, clear inhibition of the growth of O. oeni by Lb. lindenri was observed (Fig. 4.8).

G 140

(a) pH 5.5 (b) pH 3.5 10 10

8 8

6 6

4 4

2 2 Populations (log CFU/ml)

0 0 02 46 81002 4 6810 (c) pH 5.5 (d) pH 3.5 10 10

8 8

6 6

4 4

2 2 Populations (log CFU/ml)

0 0 024681002 46 810 Incubation time (day) Incubation time (day)

Fig. 4.7 Mixed culture of Oenococcus oeni 1 with either Lactobacillus kunkeei or Lactobacillus lindneri in MRST at pH 5.5 and pH 3.5, 25 C. (a), (b) Populations of O. oeni in single and mixed culture, ( ) O. oeni (single culture); ()O. oeni with Lb. kunkeei;( )O. oeni with Lb. lindneri (c), (d) Populations of Lb. kunkeei and Lb. lindneri, ()Lb. kunkeei (single culture); ()Lb. kunkeei with O. oeni;( )Lb. lindneri (single culture); ( ) Lb. lindneri with O. oeni.

ab

a b

a b

Fig. 4.8 Inhibitory effects of Lactobacillus lindneri and Lactobacillus kunkeei on a lawn culture of Oenococcus oeni. (a) Lb. lindneri; (b) Lb. kunkeei. G 141

4.4 DISCUSSION

Grapes are a primary source of microorganisms associated with winemaking (Fleet

2001). Despite the overall significance of lactic acid bacteria in wine production, there are only occasional reports of their isolation from grapes (Chapter 2). Oenococcus oeni is particularly important because of its role in conducting the malolactic fermentation

(van Vuuren and Dicks 1993; Bartowsky 2005) but, careful analysis of the literature suggests that, it has never been isolated from grapes, but possibly the leaves of grape vines (Weiller and Radler 1970). Its ecological origin in wine production remains a mystery but, like the principal wine yeast, S. cerevisiae, it probably establishes residence in the wine environment after initial introduction from grapes (Fleet et al.

2002). Little success was experienced in isolating lactic acid bacteria from wine grapes by direct agar plate culture (Chapter 3). Instead, a predominance of B. thuringiensis was found by this method, and this correlated with the widespread use of this organism as a bioinsecticide in Australian viticulture (Chapter 3, Bae et al. 2004). Consequently, the study presented in this Chapter focused on enrichment culture as an approach to determine the diversity of lactic acid bacteria associated with wine grapes, and used conditions (pH 3.5, pH 5.5, ethanol and autoenrichment) that prevented the growth of B. thuringiensis and its potential to interfere with the recovery of lactic acid bacteria (Bae et al. 2004)

Only four of the 43 batches of grapes examined gave isolates of lactic acid bacteria by direct plating, suggesting that initial populations were very low and less than 102 CFU/g.

Enrichment substantially increased the isolation frequency of lactic acid bacteria from the grapes, with damaged berries giving more isolates. Nevertheless, many samples of both undamaged and damaged berries did not give detectable lactic acid bacteria. Either

G G 142 they were absent from the grapes or they were overgrown by other species during enrichment. Surprisingly, several species of bacteria, other than lactic acid bacteria, were isolated from the enrichment cultures, including those with added ethanol or decreased pH as further selective conditions. The population of these bacteria after 5-10 days of enrichment was about 105-107 CFU/mL. The two most frequently isolated species in this category were Sporolactobacillus inulinus and Asaia siamensis.

Sporolactobacillus inulinus was first described by Kitahara and Suzuki (1963) and is an unusual endospore forming species that produces lactic acid by homofermentation. It has the potential to survive at low pH and tolerate ethanol (Sanders et al. 2003). Asaia siamensis was first described by Katsura et al. (2001) and is considered within the genera of acetic acid bacteria. This is the first time these two species have been isolated from wine grapes. Their potential significance in wine production is not known and requires investigation. It is possible that the growth of these and other bacteria during enrichment suppressed the growth of O. oeni and other lactic acid bacteria, but this requires further study. It may be necessary to design more highly selective enrichment media to detect the low populations of lactic acid bacteria that occur on wine grapes. It was thought that autoenrichment of grape homogenates might present more specific conditions for the growth of wine-related lactic acid bacteria, but this initiative gave only low isolation rates (Table 4.7).

Curtobacterium flaccumfaciens, frequently found by direct plate culture of wine grapes

(Chapter 3), was not detected in any of the enrichment cultures. As evidenced by their distinct, pigmented colony morphology, they were occasionally observed in grape homogenates that were analysed by agar plate culture prior to enrichment. Presumably, these species were not able to grow under the enrichment conditions used in this study.

G G 143

As mentioned already, a diversity of bacteria other than lactic acid bacteria targeted in this investigation, were found in enrichment cultures (Appendix 5). It was beyond the time-frame of this project to fully characterize these isolates. Future investigation of these bacteria is suggested.

PCR-DGGE was used as an additional method to detect and identify bacteria in enrichment cultures. Generally, good agreement was obtained between data obtained by

PCR-DGGE and plate culture isolation, but some inconsistencies were noted (Table 4.9).

The limitations of PCR-DGGE analysis in ecological studies have been discussed previously (Prakitchaiwattana et al. 2004) and include different affinity of the primer

DNA to template DNA for different species and the competitive influences when template DNAs are present in different relative amounts. These findings highlight the need for using a combination of different isolation and detection techniques in ecological studies. Multiple banding is another complicating factor that often arises with

PCR-DGGE analysis (Fig. 4.4). The heterogeneity of 16S rDNA due to the presence of multiple copies of the ribosomal genes, has been reported to produce several bands per species in PCR-DGGE analysis (Nübel et al. 1996; Fogel et al. 1999). Also, multiple banding patterns caused by heteroduplex bands, can be obtained during mixed-template

PCR when annealing occurs between template DNA with high sequence similarity

(Ferris and Ward 1997; Wintzingerode et al. 1997). Not all nine bands in the DGGE profiles shown in Figs. 4.4 and 4.5 corresponded to 16S rRNA molecules actually present in the samples, and these multiple bands could suggest the formation of chimeric molecules composed of two or more different strains within a species.

The diversity of lactic acid bacteria isolated in this study was substantially different to

G G 144 that reported by others. Lactobacillus plantarum and Lb. mali were the only species that corresponded with previous reports (Lafon-Lafourcade et al. 1983; Suárez et al. 1994;

Rodas et al. 2003). Lactobacillus lindneri, E. durans, Lc. lactis and Lb. kunkeei, the most frequently isolated species in this study, have not been mentioned previously as grape isolates. Consequently, particular attention was given to their identification, using both molecular and phenotypic tests (Table 4.8). The occurrence and prevalence of Lb. lindneri is noteworthy, as it has not been associated with wine production in the past.

However it has been described as a significant spoilage organism (Sakamoto and

Konings 2003) and, therefore, could have the potential to spoil wines. Lactobacillus kunkeei was recently described as a new species, and was originally isolated from wines undergoing stuck or sluggish fermentations (Edwards et al. 1998b). It is an undesirable species in wine production, and its association with wine grapes is reported here, for the first time. Both Lb. lindneri and Lb. kunkeei are antagonistic towards O. oeni

(particularly Lb. lindneri), and their frequent occurrence on grapes may explain the difficulty of isolating O. oeni from this habitat (Figs. 4.7, 4.8). Bacteriocin production by these species is probably the underlying mechanism of this inhibition (Navarro et al.

2000; Bauer et al. 2005) but this possibility requires confirmation. Such bacteriocins could have commercial application in preventing malolactic fermentation in wines where this reaction is not desired. The potential use of nisin, produced by Lactococcus lactis, for controlling malolactic fermentation has been reported (Radler 1990; Navarro et al. 2000; Bauer et al. 2003). The particularly strong inhibition of O. oeni by Lb. lindneri, warrants further investigation.

Species of Enterococcus, Weissella and Lactococcus are not previously known as wine microorganisms (Stiles and Holzapfel 1997), and probably do not grow at the higher

G G 145 ethanol concentrations in wine. Nevertheless, they were isolated from enrichment cultures containing 5% ethanol. The ethanol tolerance of these species requires more detailed investigation. Their association with the surface of plants and agricultural environments is well documented (Mundt et al. 1967; Sandine et al. 1972; Cai 1999;

Chen et al. 2005). Consequently, their isolation from the surfaces of grapes is not unexpected.

In summary, the incidence and populations of lactic acid bacteria on wine grapes were very low. Damaged grape berries showed a greater presence of these bacteria than undamaged berries. The diversity of bacterial species isolated from the grapes was greater than previously reported, and represented both lactic acid bacteria and non-lactic acid bacteria. Some of these bacteria (i.e. Lb. lindneri, Lb. kunkeei) could be detrimental to wine production. O. oeni was not found on grapes, but its recovery could be obscured by overgrowth from other species.

G G 146 CHAPTER FIVE

OCCURRENCE AND SIGNIFICANCE OF Bacillus thuringiensis ON WINE GRAPES

5.1 INTRODUCTION

Bacillus thuringiensis (commonly referred to as Bt) is well-known as a biological insecticide and has been used in agricultural practices for over 50 years (Glare and

O’Callaghan 2000; Nester et al. 2002; López et al. 2005). It is commercially available as a powdered or granulated formulation that contains a mixture of endospores and parasporal which are toxic to insects. Bacillus thuringiensis is used during the cultivation of wine grapes to control infestations by apple moth and other insects. It is applied as a spray to vines and grape berries at regular intervals from bud-burst until 1-2 weeks before grape harvest (Emmett et al. 1992).

The surface of grape berries represents a phyllospheric habitat for a diversity of fungal, yeast and bacterial species that have various impacts on the efficiency and quality of wine production (Fleet et al. 2002; Fleet 2003a). Filamentous fungi such as Botrytis cinerea, and species of Penicillium and Aspergillus are well known for their ability to spoil grapes prior to harvest (Pitt and Hocking 1997; Zahavi et al. 2000; Hocking and

Pitt 2003). Recently, the mycotoxin, ochratoxin A, has been found in some wines, and its occurrence is related to the growth of Aspergillus carbonarius and other species on grapes during cultivation (Cabañes et al. 2002; Sage et al. 2002; Serra et al. 2003;

Battilani et al. 2004). Yeast species within the genera Kloeckera (teleomorph

Hanseniaspora), Metschnikowia, Candida, Kluyveromyces and Pichia are frequently

G G 147 found on the surface of grapes, become part of the microflora of freshly extracted grape must, and contribute to the indigenous flora of many wine fermentations. They complement the action of Saccharomyces cerevisiae, the principal yeast of the alcoholic fermentation. Saccharomyces cerevisiae also occurs on grapes, but at much lower populations (often non-detectable) than other yeasts. There is an increasing interest in exploiting the contributions of the indigenous non-Saccharomyces yeasts to wine fermentations, as they have the potential to enhance the complexity and distinctiveness of wine flavour (Heard 1999; Pretorius 2000; Fleet 2001; Bartowsky et al. 2004). Many wines undergo a malolactic fermentation which is a secondary process that occurs after the main alcoholic fermentation by yeasts. Lactic acid bacteria, principally O. oeni, are responsible for this fermentation, that can enhance the acid balance, flavour, and microbial stability of wine (Henick-Kling 1993; Bartowsky and Henschke 1995; Fleet

2001; Liu 2002; Bartowsky 2005). The origin of malolactic bacteria, apart from those added as starter cultures, is not clear although the grape surface is considered to be a primary candidate (Wibowo et al. 1985; Lonvaud-Funel 1999). The use of B. thuringiensis in viticultural practices could affect the microflora associated with grapes and their contribution to wine production. The ecological surveys reported in Chapter 2 showed B. thuringiensis to be the most prevalent species on wine grapes.

Despite the application of B. thuringiensis to grapevines, its population on grapes and potential impact on microorganisms significant in wine production have not been the focus of specific investigation. In addition, recent studies implicating some strains of B. thuringiensis in food poisoning outbreaks, highlight the need to know more about the survival and growth of this species in grape juice and wine (Jackson et al. 1995;

Damgaard et al. 1996; Rivera et al. 2000). This chapter reports the populations of B.

G G 148 thuringiensis on wine grapes harvested at various stages during cultivation, the potential for its growth and survival in grape juice and wine, and its antagonistic activity towards a range of grape fungi, wine yeasts and malolactic bacteria.

5.2 MATERIALS AND METHODS

5.2.1 Microbial strains

Bacillus thuringiensis was isolated from two commercially available insecticides,

Dipel“DF (Abbott Laboratories, Chicago, USA) and Delfin“WG (Thermo Trilogy Co.,

Columbia, USA) that use this organism as the active constituent. Insecticide (1 g) was suspended in 9 mL of 0.1% Bacteriological Peptone (Oxoid, Melbourne, Australia), diluted, spread inoculated on plates of Tryptone Soya Agar (TSA, Oxoid) and incubated at 30qC for 24 h. Colonies were isolated and purified by streaking. These strains were maintained on slants of Nutrient Agar (NA, Oxoid), after incubating at 30qC for 24 h.

Two strains of Oenococcus oeni were similarly isolated by plating samples from commercial starter culture products onto MRSA agar. MRSA agar consisted of de Man,

Rogosa and Sharpe agar (MRS, Oxoid) supplemented with 15% (v/v) apple juice (Berri

Pty. Ltd., Australia) and adjusted to pH 5.5 using HCl. Plates were incubated at 25qC for

7 days. O. oeni 1 was isolated from Viniflora CH35• (CHR. Hansen Pty. Ltd.,

Australia) and O. oeni 2 was isolated from Viniflora Oenos• (CHR. Hansen Pty. Ltd.).

Other cultures of bacteria, yeasts and filamentous fungi used in the study are listed in

Table 5.2 and were obtained from collections within the University of New South Wales

(UNSW), the Research Institute (AWRI) and Food Science Australia,

CSIRO. Strains of Aureobasidium pullulans, Cryptococcus species, Metschnikowia

G G 149 pulcherrima, Rhodotorula sloofiae and Sporobolomyces roseus were originally isolated from wine grapes cultivated in the Hunter Valley region, New South Wales. They were provided by laboratory collegues and PhD students Ms. Ai Lin Beh and Ms Cheunkit

Prakitchaiwattana.

5.2.2 Grape samples

Grapes samples, collected during October - February 2001-2002 and October - February

2002-2003, were obtained from six commercial vineyards located in the upper and lower Hunter Valley, Mudgee and Griffith regions of New South Wales, Australia.

Grapes were sampled at several phenological stages of development, and these stages were referred to in Chapter 3, Table 3.2. Samples included four varieties of red grapes

(Cabernet Sauvignon, Merlot, Shiraz, Tyrian) and three varieties of white grapes

(Chardonnay, Sauvignon Blanc, Semillon). Each sample consisted of small clusters or bunches of grapes (approx. 250 g) that were aseptically removed from at least five different vines within the vineyard. Duplicate samples were taken from each vineyard.

Samples were stored at 4qC and analysed within 24 h of harvest from the vine.

5.2.3 Isolation and identification of B. thuringiensis from grapes

Individual grape berries were randomly and aseptically removed from each cluster or bunch. Samples (50 g) were suspended in 450 mL of sterile 0.1% Bacteriological

Peptone (Oxoid), containing 0.01% Tween 80 (Sigma, Castle Hill, NSW, Australia) in a conical flask and orbitally shaken at 180 rpm for 30 minutes. The rinse from these grapes was diluted in 0.1% Bacteriological Peptone (Oxoid) solution and samples (0.1 mL) of each dilution were spread inoculated onto plates of Plate Count Agar (PCA,

Oxoid) supplemented with 100 mg/L of cycloheximide (Sigma). After incubation of

G G 150 these plates at 30qC for 48 h, presumptive B. thuringiensis colonies were counted and selected. Isolates were purified by streaking and their identity as B. thuringiensis was confirmed by morphological observation (Gram positive, spore-forming rod) and sequencing of the 16S ribosomal DNA.

Extraction and PCR amplification of DNA from bacterial cultures followed the methods of Nübel et al. (1996) and Garbeva et al. (2001) using the universal primers flanking the region between 968 and 1401 (E. coli numbering). These primers were synthesized and provided by SigmaGenosys Australia Pty. Ltd. The PCR mixture contained 1u PCR buffer (10 mM Tris-HCl [pH 8.3], 50 mM KCl), 0.2 PM of each primer, 200 PM of each dNTP (Roche Diagnostics Corp., Indianapolis, IN, USA), 1.5 mM MgCl2, 1.25 U of Taq

DNA polymerase (AmpliTaq•, Roche Molecular Systems Inc., Branchburg, New

Jersey, USA) and 10 ng of extracted DNA in 50 PL of final volume. PCR amplifications were performed with a PC-960 thermal cycler (Corbett Research, NSW, Australia) with the following cycling program: initial denaturation at 94qC for 2 min, 10 cycles at 94qC for 30 sec, 50qC for 30 sec, and 72qC for 45 sec, followed by 20 times the same cycle with each successive cycle at 5 sec longer elongation time. The final elongation was conducted for 10 min at 72qC. The PCR amplicons were confirmed by 1.2% agarose gel electrophoresis and used for sequencing analysis.

Sequencing was carried out with the ABI PRISM® BigDye™ Terminators v3.1 Cycle

Sequencing Kit (Applied Biosystems Inc., Foster City, CA, USA) and assayed at the

Automated DNA analysis facility, School of Biotechnology and Biomolecular Sciences,

University of New South Wales. The sequences were compared with the non-redundant

G G 151 nucleotide database at GENBANK, using BLAST to determine the identities.

5.2.4 Isolation of B. thuringiensis from wines and grape juice

Nine samples of wine (six of Chardonnay and three of Shiraz) that had completed malolactic fermentation in 2001, and 12 samples of juice were obtained from different tanks or barrels at a commercial winery in the Hunter Valley. Wines and juice were diluted in 0.1% Bacteriological Peptone (Oxoid) solution and samples (0.1 mL) of each dilution were spread inoculated on NA containing 100 mg/L cycloheximide (Sigma), and incubated for 1 to 3 days at 30qC. Colonies were identified by morphology and sequencing of the 16S ribosomal DNA as described.

5.2.5 Interaction between B. thuringiensis and other microorganisms

Five strains of B. thuringiensis isolated from Delfin“WG, five strains of B. thuringiensis isolated from DiPel“DF and three isolates from grape samples were characterized for their interaction against species of bacteria, yeasts and fungi, listed in Table 5.1, using the spot on lawn assay (Földes et al. 2000; Addis et al. 2001) and the deferred antagonism assay (Paik et al. 1997).

For the spot on lawn assay, the appropriate medium for either fungi, yeasts or bacteria was used as the basal medium. Twenty milliliters of molten Potato Dextrose Agar (PDA,

Oxoid) for fungi, GYP agar (1% D-Glucose, 1% Yeast Extract, 1% Peptone, 2% Agar) for yeast, TSA, MRS agar and MRSA agar for bacteria were cooled to 50qC, inoculated with freshly cultured suspensions (105-106 CFU/mL) of either fungal conidia, yeast cells or bacterial cells, mixed thoroughly, and allowed to solidify. After solidification, each seeded medium was spot-inoculated with the strains of B.

G G 152 thuringiensis that had been freshly pre-cultured. For the deferred antagonism assay, fresh cultures of B. thuringiensis were spot-inoculated onto TSA and incubated overnight at 30qC. Species to be tested for interaction were inoculated at 105-106

CFU/mL into the appropriate molten agar medium (5 mL) and poured as an overlay into the plates spot inoculated with B. thuringiensis. Inoculated plates were incubated at

25qC for 4 days (fungi and yeasts) and at 25qC or 30qC for 1-7 days (bacteria).

Microbial interactions were observed as zones of inhibited growth surrounding the spot cultures of B. thuringiensis. These experiments were repeated three times.

5.2.6 Growth and survival of B. thuringiensis in grape juice and wine

Growth of B. thuringiensis was examined in red grape juice (15.5 qBrix, pH 3.37, preservative free, Berri Pty. Ltd., Australia), white grape juice (22.0 qBrix , pH

3.88, preservative free, Freedom Foods Pty. Ltd., Australia), Shiraz red wine (56 mg/L total sulphur, 10 mg/L free sulphur, 0.7 g/L glucose/fructose, 13.0% alcohol) and

Chardonnay white wine (111 mg/L total sulphur, 11 mg/L free sulphur, 3.7 g/L glucose/fructose, 12.5% alcohol). The wines were commercial products (Jacobs

Creek“, South Australia). The pH of each grape juice and wine was aseptically adjusted to values between pH 3.0 and 6.0 with 10 N NaOH and 0.2 N HCl. Strains of

B. thuringiensis grown for 20 h at 30qC in Nutrient Broth (NB, Oxoid) to 107-108

CFU/mL were inoculated (1%, v/v) into 150 mL of each juice and wine, and incubated at 25qC. Samples (1.0 mL) were aseptically removed at various intervals, for analysis of

B. thuringiensis populations by spread plating onto NA.

G G 153

5.2.7 Growth of microorganisms in mixed culture with B. thuringiensis

To investigate interactive growth between B. thuringiensis and S. cerevisiae, mixed cultures were conducted in red grape juice (Berri Pty. Ltd., Australia). Saccharomyces cerevisiae AWRI 1010, grown in GYP broth at 25qC for 20 h, and B. thuringiensis grown in NB at 30qC for 20 h, were inoculated onto 200 mL of grape juice in a 250 mL

Schott bottle (Crown Scientific, Melbourne, Australia). The culture was incubated at

25qC for 30 h. Samples were removed every 3 h to determine the populations of S. cerevisiae and B. thuringiensis by spread plating onto GYP agar supplemented with 100 mg/L oxytetracycline (Sigma) and NA supplemented with 100 mg/L cycloheximide

(Sigma), respectively. Control, single cultures of each species in the juice were also conducted.

To investigate the interactive growth between B. thuringiensis and O. oeni, mixed cultures were conducted in MRSA broth at pH 5.5 and 6.5. B. thuringiensis grown up as previously described and O. oeni grown up in MRSA broth at 25qC for 5 days were inoculated into 150 mL of MRSA broth and incubated at 25qC for 6 days. Every 6 or 12 h, samples were removed to determine the populations of O. oeni and B. thuringiensis by spread plating onto MRSA agar and NA, respectively. Control, single cultures of each species were conducted in MRSA broth.

5.3 RESULTS

5.3.1 Isolation of B. thuringiensis from commercial insecticides

The insecticides, Delfin“WG and DiPel“DF gave total viable counts of 2.7u1010 CFU/g

G G 154 and 1.9u1010 CFU/g, respectively. Colonies on plates of NA had similar appearance and were identified as B. thuringiensis by their morphology and sequencing of the 16S ribosomal DNA.

5.3.2 Occurrence of B. thuringiensis on grapes, juice and wine samples

Fig. 5.1 shows the populations of B. thuringiensis on several grape varieties sampled during cultivation from six commercial vineyards for the 2002-2003 season. Vineyards

A (Upper Hunter Valley) and C (Lower Hunter Valley) were also examined throughout the 2001-2002 season. The insecticides containing B. thuringiensis were applied four times for Vineyards A and C and only up until veraison for Vineyards D, E, and F

(arrows Fig. 5.1).

Bacillus thuringiensis was consistently isolated from all grape varieties in vineyards A and C throughout the period of grape cultivation over two consecutive vintages, 2001-

2002 and 2002-2003. Its population varied between 102-106 CFU/g, with higher populations being found in the early stages of grape cultivation. There was no obvious influence of grape variety on the presence of B. thuringiensis, but some vineyards tended to give grapes with higher populations. At the time of harvest, B. thuringiensis populations on grapes varied between 102-104 CFU/g for vineyards A and C (Fig. 5.1a- d). As determined by culture on PCA, B. thuringiensis was the most prevalent bacterial species present on grapes. The populations of B. thuringiensis on grapes from vineyards

D, E, and F were not detectable (<50 CFU/g) after veraison, but prior to this time, there were higher initial populations (Fig. 5.1e, f).

Bacillus thuringiensis was inconsistently isolated from wines at different stages of

G 155

vinification in a large winery in the Hunter Valley (Table 5.1). It was found at 5x101 CFU/mL in two out of four Traminer/Chardonnay juice samples before fermentation and in six out of eight fermenting Chardonnay samples at 5x1013 -5x10 CFU/mL. It was not isolated (< 50 CFU/mL) from nine samples of Shiraz or Chardonnay wines that had completed alcoholic and malolactic fermentations and were at the stage of bulk storage in tanks or barrels.

(a) Vineyard A, 2001-2002 (b) Vineyard C, 2001-2002 7

6 scs

5 c sm sm 4 s

3

2

1

0 0VI0II III IIIa IV I II III IIIa IV V (c) Vineyard A, 2002-2003 (d) Vineyard C, 2002-2003 7

6 ssm sm 5

4

3 s c Log CFU/g Log CFU/g 2 cs

1

0 I II III IV V I II III IV V (e) Vineyards D and E, 2002-2003 (f) Vineyard F, 2002-2003 7

6 m c 5 s smsb s cs 4 t

3 sb cs Log CFU/g 2 c

1

0 I II III IV V I II III IV V Phenological stage of berry development Phenological stage of berry development

Fig. 5.1 Populations of Bacillus thuringiensis on grapes during cultivation in different regions of New South Wales, Australia. (a) Vineyard A, 2001-2002; (b) Vineyard C, 2001-2002; (c) Vineyard A, 2002-2003; (d) Vineyard C, 2002-2003; (e)Vineyards D and E, 2002-2003; (f) Vineyard F, 2002-2003. Phenological stages of berry development are given in Table 3.2. Arrows indicate stage where B. thuringiensis was applied. Shiraz (s, ) ; Cabernet Sauvignon (cs, ); Chardonnay(c, ); Semillon (sm, ); Tyrian (t, ); Merlot (m, ); Sauvignon Blanc (sb, ). G 156

Table 5.1 Population of Bacillus thuringiensis in wines during vinification

Fermentation stage Wine Total population (CFU/mL) Must before fermentation Traminer 3.5u101 Chardonnay 1.5u101 Chardonnay ND Chardonnay ND Lees ferment Chardonnay 3.8u103 Natural alcoholic fermentation Chardonnay ND Chardonnay 1.8u102 Chardonnay 3.5u101 Chardonnay 2.6u102 Chardonnay 1.5u101 Chardonnay 5.5u101 Chardonnay ND Induced alcoholic fermentation Chardonnay 4.3u103 Conservation Shiraz ND Shiraz ND Shiraz ND Chardonnay ND Chardonnay ND Chardonnay ND Chardonnay ND Chardonnay ND Chardonnay ND ND, Not detected (<50CFU/mL)

5.3.3 Interaction between B. thuringiensis and microorganisms significant in wine production

5.3.3.1. Yeasts

As determined by the spot on lawn test, none of the yeasts examined were inhibited by

B. thuringiensis (Table 5.2). However, with the deferred overlay test, there was inhibition of some yeasts, namely, species of Cryptococcus, Debaryomyces,

Metschnikowia, Pichia, Rhodotorula and Sporobolomyces. However, the principal wine yeast, S. cerevisiae was not inhibited by B. thuringiensis (Fig. 5.2a).

5.3.3.2. Bacteria

Bacillus thuringiensis was antagonistic towards other Bacillus species such as B. cereus, B. mycoides and B. subtilis (Table 5.2). Clear inhibition of the growth

G 157

principal malolactic bacterium, was observed by the deferred overlay test (Fig. 5.2b). In

addition, Lactobacillus brevis and Pediococcus pentosaceus were slightly inhibited by B.

thuringiensis. The spot on lawn assay could not be performed with lactic acid bacteria,

because the pH of the basal agar medium became acidified by the growth of these

bacteria. As a consequence, B. thuringiensis did not grow as a spot culture. Inhibitory

activity of B. thuringiensis was not detected against representative Gram-negative

phytopathogenic organisms such as Curtobacterium flaccumfaciens, Erwinia

carotovora, Pseudomonas syringae, and Xanthomonas campestris.

5.3.3.3. Filamentous fungi

Bacillus thuringiensis gave clear inhibition of the growth of Botrytis cinerea, Alternaria

infectoria, Aspergillus aculeatus, Aspergillus carbonarius, and Cladosporium

cladosporioides by both assay methods (Table 5.2, Fig. 5.2c, d).

ab

c d

10mm

Fig. 5.2 Inhibitory effects of Bacillus thuringiensis on microorganisms of significance in wine production. (a) B. thuringiensis (spot) on a lawn culture of Saccharomyces cerevisiae PDM; (b) B. thuringiensis (spot) on a lawn culture of Oenococcus oeni 1; (c) B. thuringiensis (spot) on a lawn culture of Botrytis cinerea FRR 5215; (d) B. thuringiensis (spot) on a lawn culture of Aspergillus carbonarius FRR 5374. Each plate had representative B. thuringiensis strains isolated from DelfinR WG, DiPelR DF and grapes 158

Table 5.2 Inhibitory activity of Bacillus thuringiensis against microorganisms of significance in wine production as determined by spot on lawn and deferred culture assays

Assay a Assay Microorganism tested Microorganism tested SD SD

Yeasts Saccharomyces cerevisiae SWd - - d Aureobasidium pullulans - - Saccharomyces cerevisiae UCD522 - - Aureobasidium pullulans - - Saccharomyces cerevisiae - - Aureobasidium pullulans - - Sporobolomyces roseus - + Aureobasidium pullulans - - Zygosaccharomyces bailli - - Aureobasidium pullulans - - Fungi Candida lipolytica - - Alternaria infectoria FRR 4821 + + Candida stellata - - Aspergillus aculeatus FRR 5376 + + Cryptococcus chernovii - - Aspergillus carbonarius FRR 5374 + + Cryptococcus laurentii - - Aspergillus niger FRR 5375 - - Cryptococcus magnus - + Botrytis cinerea FRR 5215 + + Debaryomyces hansenii - + Cladosporium cladosporioides FRR 4778 + + b Kloeckera apiculata AWRI 867 - - Penicillium crustosum FRR 4775 +/- +/- c Kloeckera apiculata FRR 610 - - Penicillium expansum FRR 4776 +/- +/- Kluyveromyces marxianus FRR 1586 - - Bacteria Metschnikowia pulcherrima - - Bacillus cereus UNSWe 052300 + + Metschnikowia pulcherrima - +/- Bacillus cereus UNSW 055600 + + Metschnikowia pulcherrima - +/- Bacillus mycoides UNSW 001200 + + Metschnikowia pulcherrima - +/- Bacillus subtilis UNSW 030702 + + Curtobacterium flaccumfaciens - Pichia anomala FRR 2169 - + - Erwinia carotovora UNSW 031700 - - Pichia membranifaciens - + Gluconobacter oxydans UNSW 030300 - - Rhodotorula sloofiae - + Lactobacillus brevis UNSW 055100 NA +/- Saccharomyces cerevisiae AWRI 1010 - - Lactobacillus casei UNSW 060400 NA - Saccharomyces cerevisiae AWRI 350d - - Lactobacillus plantarum UNSW 084800 NA - Saccharomyces cerevisiae AWRI 796d - - Leuconostoc mesenteroides UNSW 060700 NA - d Saccharomyces cerevisiae AWRI R2 - - Oenococcus oeni 1 NA + d Saccharomyces cerevisiae Cru-Blanc - - Oenococcus oeni 2 NA + d Saccharomyces cerevisiae PDM - - Pediococcus pentosaceus USNW 047200 NA +/- Saccharomyces cerevisiae Primeur d - - Pseudomonas syringae UNSW 036900 - - Saccharomyces cerevisiae Sauvignon L3 d - - Xanthomonas campestris UNSW 031500 - - -, no inhibition; +, clear zone of inhibition; +/-, zone of inhibition was hazy, not clear; NA , not tested a S, Spot on the lawn assay; D, Deferred culture assay bAWRI, Australian Wine Reserach Institute, Adelaide c FRR, Food Research Laboratory, Food Science Australia, CSIRO, North Ryde d Commercial yeast starter culture of MaurivinR (Maurivin Yeast Australian Pty. Ltd.) and CanadivinTM (Burns Philip and Co.) e UNSW, School of Biotechnology and Biomolecular sciences, The University of New South Wales, Sydney All other cultures from Food Science and Technology, UNSW, Sydney; each culture represents a different isolate/strain 159

5.3.4 Survival and growth of B. thuringiensis in grape juice and wine

Bacillus thuringiensis did not grow in samples of white and red grape juice or in samples of white or red wine (Fig. 5.3). However, viable cells (103 -104 CFU/mL) inoculated into these samples remained stable and did not die off throughout the 12 day incubation period. This behavior was observed for juice or wine samples adjusted to either pH 3.0, 3.5 or 4.0 (Fig. 5.3) and pH 4.5, 5.0 or 6.0 (data not shown).

5 red juiceA red juice B 4

3

2 Log CFU/mL 1

0 5 white juiceAB white juice 4

3

2 Log CFU/mL 1

0 5 red wineA red wine B 4

3 CFU/mL 2 Log 1

0

5 white wineA white wine B 4 3

2 Log CFU/mL 1

0 0 24 68 10 12 0264 8 10 12

Incubation time (days) Incubation time (days)

Fig. 5.3 Survival and growth of Bacillus thuringiensis in grape juice and wine. (A) B. thuringiensis from DelfinR WG; (B) B. thuringiensis from DiPelR DF. pH 3.0 ( ), pH 3.5 ( ), pH 4.0 ( ). 160

5.3.5 Interactive growth of B. thuringiensis and S. cerevisiae

The growth of S. cerevisiae in grape juice was not affected by the presence of B.

thuringiensis at initial populations of 103 -104 CFU/mL (Fig. 5.4). The growth profile of

S. cerevisiae in the presence of either strain of B. thuringiensis was similar to that of the

control (Fig. 5.4a). Bacillus thuringiensis did not grow in the juice, but its population

remained stable throughout fermentation with S. cerevisiae (Fig. 5.4b).

9 (a)

8

7 Log CFU/mL 6

5 5 (b)

4

3 Log CFU/mL 2

1 0 36912151821242730 Incubation time (hours)

Fig. 5.4 Mixed culture of Bacillus thuringiensis and Saccharomyces cerevisiae AWRI 1010 in red grape juice at 25 C. (A) Population of S. cerevisiae in mixed culture S. cerevisiae with B. thuringiensisWG ( ); S. cerevisiae with B. thuringiensis DF ( ); S. cerevisiae (single culture, control) ( ); (B) Population of B. thuringiensis in mixed culture; B. thuringiensis WG with S. cerevisiae ();B. thuringiensis WG (single culture, control) ( ); B. thuringiensis DF with S. cerevisiae ();B. thuringiensis DF (single culture, control) ( ). G 161

5.3.6 Interactive growth of B. thuringiensis and O. oeni

Oenococcus oeni is nutritionally fastidious and can be problematic to culture in liquid media. However, most strains of this species grow in MRS broth supplemented with either tomato juice or apple juice (Pan et al. 1982; Wibowo et al. 1985; Fugelsang 1997).

Bacillus thuringiensis also exhibited good growth in MRSA, so it was chosen as the medium in which to examine the interactive growth of the two species. Both strains of

O. oeni behaved similarly in these growth experiments. The growth profile of O. oeni 1 and 2 in MRSA at pH 5.5 was not affected by the presence of B. thuringiensis (Fig. 5.5a, c). As single cultures at pH 5.5, B. thuringiensis exhibited a tendency to die off during the first few days, and then commenced growth (Fig. 5.5b, d). However, in mixed culture with O. oeni, its slow, initial loss of viability continued.

The growth of O. oeni in MRSA at pH 6.5 was enhanced when it was cultured in combination with B. thuringiensis (Fig. 5.6a, c). Final populations of O. oeni were about

5-10 times greater (108-109 CFU/mL) when grown in the presence of either strain of B. thuringiensis. After a short lag phase, both strains of B. thuringiensis, grew in MRSA, pH 6.5, to 107–108 CFU/mL by day 3. This behavior was observed for B. thuringiensis grown singly and in mixed culture with O. oeni (Fig. 5.6b, d). However, for the mixed cultures, but not single cultures, there was a sharp loss in the viability of B. thuringiensis between days 3–4. This corresponded to a stage when O. oeni had reached maximum populations of 109-1010 CFU/mL (Fig. 5.6a, c).

G 162

10 8 (a) O. oeni 1 (b) B. thuringiensis + O. oeni 1 9 7

8 6

7 5 Log CFU/mL

6 4

5 3 10 8 (c) O. oeni 2 (d) B. thuringiensis + O. oeni 2 9 7

8 6

7 5 Log CFU/mL

6 4

5 3 01234560123456 Incubation time (days) Incubation time (days)

Fig. 5.5 Mixed culture of Bacillus thuringiensis and Oenococcus oeni in MRSA at pH 5.5, 25 C. (a) Population of O. oeni 1; (b) Population of B. thuringiensis; (c) Population of O. oeni 2; (d) Population of B. thuringiensis. O. oeni with B. thuringiensis WG O. oeni with B. thuringiensis DF O. oeni (control, single culture) B. thuringiensis WG with O. oeni B. thuringiensis WG (control, single culture) B. thuringiensis DF with O. oeni B. thuringiensis DF(control, single culture) 163

10 9 (a) O. oeni 1 (b) B. thuringiensis + O. oeni 1 8 9 7

8 6

7 5 Log CFU/mL 4 6 3

5 2 10 9 (c) O. oeni 2 (d) B. thuringiensis + O. oeni 2 8 9 7

8 6

7 5

Log CFU/mL 4 6 3

5 2 01234560123456 Incubation time (days) Incubation time (days)

Fig. 5.6 Mixed culture of Bacillus thuringiensis and Oenococcus oeni in MRSA at pH 6.5, 25 C. (A) Population of O. oeni 1; (B) Population of B. thuringiensis; (C) Population of O. oeni 2; (D) Population of B. thuringiensis. O. oeni with B. thuringiensis WG O. oeni with B. thuringiensis DF O. oeni (control, single culture) B. thuringiensis WG with O. oeni B. thuringiensis WG (control, single culture) B. thuringiensis DF with O. oeni B. thuringiensis DF(control, single culture) G 164

5.4 DISCUSSION

The surface of grape berries is a phyllospheric habitat and, like other phyllospheres, there can be dynamic interaction between the resident microflora (Belanger and Avis

2002; Fleet 2003a). These interactions will determine the overall microbial ecology of the grape at the time of harvest and the microbial populations that transfer to the grape juice and wine production. The bacterial ecology of wine grapes has been a neglected topic of research in the fields of enology and viticulture. It is generally accepted that grapes are an important source of lactic acid bacteria and acetic acid bacteria associated with wine production (Lafon-Lafourcade et al.1983; Wibowo et al. 1985; Drysdale and

Fleet 1988; Lonvaud-Funel 1999) but the analytical evidence supporting this view is very limited. The presence of B. thuringiensis on wine grapes, as consistently found in this study (Chapter 3), has not been previously reported although there is a brief mention of its isolation from table grapes and other fruits, where its occurrence was linked to its use as a biopesticide during fruit cultivation (Bidochka et al. 1987). It is not possible to conclude from this study if B. thuringiensis occurred on grapes as spores or as vegetative cells although it is reported that commercial Bt pesticides usually contain a mixture of spores, endotoxin crystals, vegetative cells, cell debris and carry over of material from the fermentation (Glare and O’Callaghan 2000). There is an abundance of spores in the pesticide preparations and there will be an opportunity for these to germinate during reconstitution and storage of the pesticide solution and, possibly, on the grape surface.

The population of B. thuringiensis on grapes ranged between 100-106 CFU/g but this variation could not be correlated with any particular intrinsic or extrinsic factor such as

G G 165 grape variety or timing of application. However, populations were generally higher on grapes at the early stage of cultivation when they were smaller and not bunched and, presumably, when the spray had greater access to individual berries. Many factors are likely to affect the population of B. thuringiensis on grapes and these include the particular commercial preparation of B. thuringiensis, spraying technique, acidity of the grape surface, and environmental factors such as UV radiation, temperature, humidity and rainfall (Griego and Spence 1978; Leong et al. 1980; Ignoffo 1992). Ignoffo et al.

(1974) found that 65% of insecticidal activity and 90% of spore viability were lost the first day after application, although some insecticidal activity remained for 7 days.

Petras and Casida (1985) reported that spore numbers declined by one order of magnitude after 2 weeks but then remained constant for 8 months following application

In viticultural practices, prolonged and successive programs of application, broad aerial dispersal and turning over of spores from soil by wind and machinery are factors which would contribute to the presence of B. thuringiensis on grapes until harvest.

Bacillus thuringiensis on grapes carries over into wine production as demonstrated by its isolation from several samples of grape juice before and during alcoholic fermentation. Inconsistent isolation of B. thuringiensis at this stage might be linked to the amount of used during juice processing, but this possibility requires investigation. The occurrence of B. thuringiensis in the winery environment raised questions about its ability to grow in grape juice and wine, and its potential to compete with the growth of S. cerevisiae during alcoholic fermentation and O. oeni during malolactic fermentation.

As mentioned already, some strains of B. thuringiensis have been linked to outbreaks of gastroenteritis (Jackson et al. 1995; Damgaard et al. 1996) and, taxonomically, Bacillus

G G 166 thuringiensis is closely related to Bacillus cereus, a well known food borne pathogen that causes emetic and diarrhoeal forms of food poisoning (Granum 1994; Rusul and

Yaacob 1995; Granum 2001). In addition, it was recently shown that the distribution of enterotoxin genes is ubiquitous in B. thuringiensis and that its toxicity is equivalent to the toxicity associated with B. cereus isolated from cases of diarrhoeal food poisoning

(Rivera et al. 2000). The infectious or toxigenic dose of B. cereus is greater than 105 cells/g (Jensen and Moir 2003) and, by analogy, should be similar for B. thuringiensis.

However, such populations were not found on grapes at the time of harvest. Moreover,

B. thuringiensis was not able to grow in grape juice or wine, even when the pH was increased to values as high as 6.0. Consequently, there would be no public health or spoilage risks from the presence of B. thuringiensis in grape juice or wine. More general studies have shown that B. thuringiensis and the agricultural use of Bt products have an excellent safety record (Glare and O’Callaghan 2000; Siegel 2001; Nester et al. 2002).

Although B. thuringiensis has the potential to inhibit some yeasts species frequently associated with wine grapes, it was not active against several wine strains of

S. cerevisiae (Table 5.1). Consequently, it is unlikely that the routine application of Bt products to grape vines accounts for the inability of many researchers to isolate

S. cerevisiae from wine grapes (Fleet et al. 2002). Furthermore, B. thuringiensis did not adversely affect the growth of S. cerevisiae in grape juice (Fig. 5.4). Thus, its carry over from grapes into the juice is unlikely to impact on the conduct of the alcoholic fermentation and be a potential cause of stuck or sluggish fermentation (Bisson 1999).

The interaction of B. thuringiensis with the malolactic bacterium, O. oeni, gave inconsistent, but interesting data, depending on the assay. The plate assay showed clear

G G 167 inhibition of O. oeni by B. thuringiensis. In these assays, B. thuringiensis had grown in the medium before O. oeni and presumably produced substances that were antagonistic to O. oeni. B. thuringiensis is known to produce bacteriocins (Favret and Yousten 1989;

Paik et al. 1997) which could account for this observation. In mixed culture in liquid, however, the growth of O. oeni was not adversely affected by the presence or growth of

B. thuringiensis (Figs. 5.5, 5.6). In fact, there was a slight stimulation of the growth of

O. oeni, which was observed for both strains of O. oeni and both strains of

B. thuringiensis. Interestingly, growth of O. oeni caused loss of viability of

B. thuringiensis, especially at pH 6.5 (Fig. 5.6b, d) and presumably this may be associated with inhibitory substance(s) produced by O. oeni (Edwards et al. 1994). This interaction might account for the inability to isolate B. thuringiensis from all samples of wine that had completed malolactic fermentation. Further research is suggested to understand the mechanisms of these interesting interactions.

As demonstrated by the plate assay, B. thuringiensis has the potential to inhibit the growth of several filamentous fungi associated with the spoilage of grapes (B. cinerea) and ochratoxin A production on grapes (A. carbonarius). It remains to be determined if such interactions occur in situ at the grape surface. Current use of B. thuringiensis in viticulture to control insects could, therefore, inadvertently and unknowingly control spoilage and mycotoxigenic fungi. Field studies need to be conducted to correlate the use of B. thuringiensis in viticulture with the occurrence of spoilage and mycotoxigenic fungi on grapes.

In conclusion, data presented on this Chapter and Chapter 3 have shown, for the first time, that B. thuringiensis is a prevalent organism on wine grapes where it has the

G G 168 potential to influence the phyllospheric microflora and microbial species entering the winery environment. Although B. thuringiensis carries over into the winery environment, it is not able to grow in grape juice or wines and, does not influence the growth of S. cerevisiae or O. oeni therefore, does not adversely impact on wine quality or safety.

G G 169 CHAPTER SIX CONCLUSIONS

This thesis has reported a systematic investigation of the bacterial species associated with wine grapes cultivated in several Australian vineyards. Grape samples were examined as they matured throughout cultivation. A combination of cultural and molecular methods was used to analyse the bacterial species associated with these grapes. Three red grape varieties (Shiraz, Cabernet Sauvignon, Merlot) and three white grape varieties (Chardonnay, Semillon, Sauvignon Blanc) were examined, and analyses were conducted over the 2001-2002, 2002-2003 and 2003-2004 seasons. Various factors likely to affect the bacterial ecology of grapes were considered. These factors included vineyard location, climatic influence, pesticide applications, and grape berry damage.

The total bacterial population of mature undamaged grapes at the time of commercial harvest was consistently low at 102-103 CFU/g. Higher populations (103-106 CFU/g) were found on grapes at earlier stages of maturity and correlated with application of the biological pesticide Bt which is a commercial preparation of the bacterium, B. thuringiensis. Bacillus thuringiensis was the most prevalent bacterial species found on wine grapes, as determined by plate culture, enrichment and PCR-DGGE methods. This was a novel finding in the field of wine microbiology, but not unexpected considering the widespread use of Bt as a biological pesticide in Australian viticulture.

After B. thuringiensis, C. flaccumfaciens was the next most prevalent bacterial species detected on wine grapes, although not consistently found on all grapes. Its presence was found by both plate culture and PCR-DGGE methods. Its population was low and rarely

G G 170 exceeded 103-104 CFU/g. Curtobacterium flaccumfaciens is a well known phyllospheric organism, so its occurrence on the surface of grapes would not unexpected. The oenological significance of its presence on wine grapes needs further investigation.

A diversity of other bacterial species (e.g. Pseudomonas, Sphingomonas, Ralstonia,

Arthrobacter, Microbacterium) was found on grapes by either culture or PCR-DGGE methods, but these occurrences were sporadic, at very low populations and without any viticultural correlation. Notably, Bacillus benzoevorans was consistently detected on grapes in several vineyards (B, D, F) throughout cultivation. However, it was not found by cultural methods and was only detected by PCR-DGGE analysis. Conditions for the culture of this species need to be researched. Any potential oenological significance of its association with grapes requires further study.

Lactic acid bacteria and acetic acid bacteria were rarely detected on undamaged wine grapes throughout the 2001-2002 and 2002-2003 seasons by any analytical method, suggesting that the surface of healthy, undamaged grapes is not a consistent or natural habitat for these species. Notably, the malolactic bacterium, O. oeni, was not detected on any sample of grapes, raising questions as to the origin of this important wine microorganism. Lactobacillus plantarum, noted in previous literature as being associated with wine grapes, was found on only two occasions.

Occasionally, there were discrepancies between plate culture and PCR-DGGE analysis for the detection of bacteria on wine grapes, highlighting the need for using a combination of culture and culture-independent methods for studying the microbial ecology of natural habitats, such as grapes. It is likely that populations of VBNC

G G 171 bacteria occur on the surface of grapes and more specific studies are required on this phenomenon.

Most analyses of grapes were conducted during 2002-2003. Unfortunately, this was an atypical hot and dry vintage across all vineyards examined. These conditions probably accounted for the very low populations of bacteria found on the grapes. This factor, combined with the overwhelming presence of B. thuringiensis, prevented meaningful comparisons of data to determine influences, such as vineyard location, grape variety, grape maturity, climate and viticultural practices, on the bacterial ecology of grapes.

More systematic and controlled studies of these variables are required. However, damaged grape berries generally exhibited higher bacterial populations and species diversity than mature undamaged grape berries. There was a higher incidence of wine lactic acid bacteria (e.g. Lactobacillus kunkeei) and wine acetic acid bacteria (e.g.

Gluconobacter and Acetobacter spp.) on damaged grapes. However, damaged grapes did not show the very high bacterial populations (e.g. ~106 CFU/g or more) reported in the literature and this could relate to the dry, low rainfall conditions during cultivation of grapes. Grapes experience different types of damage and further research is needed to investigate this variable.

An array of enrichment culture conditions using MRS broth, MRS + ethanol, MRST

(pH 5.5), MRST (pH 3.5) and autoenrichment in grape must was used during the 2003-

2004 vintage to specifically target the detection and isolation of lactic acid bacteria from wine grapes. In addition to plate culture, PCR-DGGE analysis was used to detect the bacteria in enrichment cultures. A greater incidence of lactic acid bacteria on wine grapes was detected by these enrichment procedures, but many samples of both

G G 172 damaged and undamaged berries did not give detectable lactic acid bacteria, reinforcing earlier conclusions about their inconsistent and low presence on wine grapes. However, damaged berries gave a much higher incidence than undamaged grapes. Lb. plantarum,

Lb. mali, Lb. kunkeei and Lb. lindneri were the main species found. The presence of Lb. plantarum and Lb. mali corresponds with previous literature but discovery of Lb. kunkeei and Lb. lindneri on wine grapes is a novel finding. Oenococcus oeni was not isolated from any grape samples by the diversity of enrichment conditions used, again, raising questions about the origin of this species in wine making. Both Lb. kunkeei and

Lb. lindneri were antagonistic to O. oeni in plate and mixed culture assays. The antagonistic activity of these bacteria and other microorganisms to O. oeni, could explain the failure to detect this species on grapes. Further research is needed on this question. Moreover, it may be necessary to develop more selective and specific enrichment conditions to detect O. oeni on grapes. It is noteworthy that the enrichment cultures used also revealed a diversity of non-lactic acid bacteria on wine grapes – especially Sporolactobacillus inulinus and Asaia siamensis. Further studies are needed to evaluate the oenological significance of these species because they have not been reported, as grape microflora in previous literature.

Because of the prevalence of B. thuringiensis on wine grapes, further studies were conducted to determine aspects of its oenological significance. Although this organism carried over from the grapes into freshly extracted juice or must, it exhibited no tendency to grow in juice or wine at pH 3.0-6.0. Its presence in juice did not adversely affect the growth of S. cerevisiae or O. oeni. Consequently, it would not contribute to sluggish or stuck fermentations. In this context, it is unlikely to adversely impact on wine quality or safety. In agar plate culture assays, however, B. thuringiensis did show

G G 173 an ability to inhibit O. oeni, and this discrepancy with liquid culture observation needs to be understood. In agar plate culture assays, B. thuringiensis also inhibited important grape spoilage (B. cinerea) and mycotoxin producing (Aspergillus spp.) fungi. These findings are putatively, oenologically very important, and emphasize the need for further research to understand the impact of this biological pesticide on the microbial ecology of grapes.

Overall, this investigation has given much new information about the bacteria associated with wine grapes and has revealed many new directions for future research.

G G 174 CHAPTER SEVEN

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APPENDIX 1 Ribosomal DNA sequence identification of bacterial isolates obtained from wine grapes by plate culture and enrichment culture

Name Accession Sequence Samplesa Number similarity (Vineyard b, Maturity stage c) (%) Acidovorax sp. MN 33..2 AJ555476.1 99 Sz (A, I) Acinetobacter sp. Hop10 AF015300.1 99 Cab (A, I) Acinetobacter sp. HJ2 AY237406.1 98 Cab (E, I) Acinetobacter sp. phenon2 AJ275041.2 98 Sab (E, I) Arthrobacter chlorophenolicus strain AY167845.1 97 Cha (B, II) SAFR-044 Arthrobacter gandensis type strain AJ316140.1 93 Cha (D, III), Cha (B, II) R5812T

Arthrobacter luteolus AJ243422.1 97 Sz (A, I) Arthrobacter oxydans X83408 95 Cha (F, II), Cha (F, IV) Arthrobacter polychromogenes X80741 92 Cha (F, V), Cha (D, I) Arthrobacter polychromogenes X80741.1 92 Sz (F, V), Sem (A, III) Arthrobacter sp. AB017650.1 92 Sz (D, II), Cha (A, V) Arthrobacter sp. Fa21 AY131225.1 91 Mer (E, II), Sab (E, II) Arthrobacter sp. CT5 AY278872.1 94 Sz (F, II), Cha (D, V), Sz (F, IV) Arthrobacter sp. JCM1339 AB070602.1 98 Sz (D, III) Arthrobacter sp. LCSAOTU6 AF506065.1 92 Cha (D, IV), Mer (E, III) Arthrobacter sp. M4 AY177360.2 88 Cab (E, III) Bacillus fumarioli AJ250059.1 97 Cha (D, IV) Bacillus megaterium strain SAFR-011 AY167865.1 97 Cab (E, III), Cha (F, IV), Sz (F, III), Sz (F, IV), Sz (A, V), Szd (F, V), Sz (A, I), Cha (A, I), Cab (E, I), Sem (C, II), Mer (E, II), Sz (F, I), Cha (F, I) Bacillus pumilus strain WN691 AY260864.1 94 Sab (E, V) Bacillus pumilus KL-052 AY030317.1 99 Sz (A, II) Bacillus senegalensis strain RS8 AF519468.1 94 Cha (F, IV), Sz (F, V) Bacillus sp. AJ297719.1 93 Sz (B, II) Bacillus spaericus 205y AF435435.1 99 Cab (A, II), Sz (D, III) Brachybacterium paraconglomeratum AJ415377.1 94 Cab (A, II) Brevibacterium acetylicum NCIMB 9889 X70313.1 98 Sz (A, I), Cab (A, I), Sem (A, I), Cha (D, III), Mer (E, III), Cha (F, I) Bacillus thuringiensis AY167823.1 96-99 AF155955 Curtobacterium fangii AY273209.1 93 Cha (D, V) Curtobacterium flaccumfaciens pv. AY273208.1 94-100 Cha (D, III), Cha (D, V), beticola Sem (A, IV), Cha (A, I), Sz (B, III), Sz (A1, V), Cab (A1, III), Cab (A1, V), Cha (A1, III), Cha (A1, IV), Cha (A1, V), Sz (A, III), Sem (C, III), Cha (B, III), Cha (A, V), Cha (F, II), Cha (F, III), Cha (F, IV), Sab (E, I), Cab (A, I), Cha (F, V), Chad (D, V)

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Name Accession Sequence Samplesa Number similarity (Vineyard b, Maturity stage c) (%) Curtobacterium flaccumfaciens strain AY167859.1 96-99 Sz (A1, II), Sem (A1, V), Sz SAFR-001 (B, IV), Cab (E, I), Sab (E, IV), Cab (E,V), Mer (E, V), Sab (E, III), Cha (A, I), Sz (D, IV), Sz (F, V) Curtobacterium sp. T2-21 AY278888.1 96-98 Cha (D, I)

Enterococcus faecalis V583 AF515223.1 98 Cab (A, II) Enterococcus mundtii AF061013.1 97 Cha (B, IV), Cha (D, III), Cha (D, IV) Enterococcus mundtii AB066266.1 98 Mer (E, V), Cha (F, III), Cha (F, IV), Cha (F, V) Erwinia cypripedii AJ233413.1 97 Cab (A, I) Erwinia rhapontici U80206.1 97 Sz (D, II) Flavobacterium sp. A230767 96 Cha (A, I) Frigoribacterium sp. 312 AF157480.1 97 Cha (D, III), Sab (E, I) Gluconobacter oxydans DQ117918 99 Sz (A1, V), Cha (A1, V), Semd (A1, V) Hafnia alvei M59155.1 97 Cha (A, I) Lactobacillus plantarum strain AJ306297.1 97 Sz (C, I) DSM1066T Lactobacillus plantarum strain WCFS1, AL935261.1 97 Sem (C, I) Lactococcus lactis subsp. lactis... AF493058.1 99 Sz (A, I) Leifsonia poae AF116342 95 Sz (F, III) Microbacterium paraoxydans AJ491806.1 94 Cha (A, I), Mer (E, III), Sab (E, I) Microbacterium phyllosphaerae strain AY167852.1 99 Sem (A, I), Cab (E, I) SAFR012 Microbacterium sp. AJ296094.1 96 Sem (A1, III), Cab (E, I), Mer (E, I) Microbacterium sp. ASD AY040877.1 99 Cha (F, V), Cab (A1, II) Microbacterium sp. MAS133 AJ251194 98 Cha (D, I) Microbacterium sp. T2-22 AY278889.1 92 Sem (A1, II), Cab (E, IV) Mer (E, III) Microbacterium terregens Y17239.1 97 Cha (D, I) Microbacterium.barkeri X77446.1 96 Sab (E, III) Micrococcus luteus AJ31275.1 98 Sz (F, V) Oxalobacteraceae bacterium MHW73 AJ556800.1 94 Sz (D, III) Pantoea agglomerans AF157694.1 99 Sz (D, I), Cab (E, II), Sz (F, I) Pantoea agglomerans AJ233423.1 91 Sz (D, II), Sz (D, III) Mer (E, II), Cha (B, III) Pantoea agglomerans strain 732 AY092079.1 98 Sz (F, II) Pantoea agglormerans AF130917.1 98 Cha (D, III) Pantoea ananatis LMG20104 AF364846.1 97 Sz (A, I), Sz (D, V), Cha (D, I), Sab (E, I), Sab (E, III) Pantoea sp. B232 AF451269.1 97 Cab (A, I), Mer (E, I), Pantoea sp. P101 AF394538.1 97 Sem (A1, V)

Pantoea stewartii AF373198.1 98 Cab (E, I) Pantoea toletana strain A64 AF130963 96 Cha (F, II) Pseudomonas aureofaciens AF301220.1 97 Cab (A, I) Pseudomonas sp. Fa20 AY131224.1 98 Sz (D, I) Pseudomonas sp. M13 AY177356.2 94 Cha (A, V), Sz (B, V)

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Name Accession Sequence Samplesa Number similarity (Vineyard b, Maturity stage c) (%) Pseudomonas sp. Sa03 AF511508.1 99 Mer (E, I) Pseudomonas sp. TB37 AY278935.1 99 Sab (E, I), Cha (F, I) Rathayibacter tritici strain SAFR-009 AY67853.1 92 Sz (D, II), Cha (D, II), Cha (D, III) Rhodococcus equi strain DSM 20307T AF490539.1 97 Cab (E, I), Cha (D, I) Rhodococcus sp. I7 AY177354.2 94 Cab (E, III), Cab (E, IV), Sz (B, III), Mer (E, III) Serratia liquefaciens AY253924.1 100 Sz (A, I) Serratia marcescens DQ144501 100 Sem (C, II) Sphingomonas parapaucimobilis D84525.1 95 Sz (D, V) Sphingomonas sp. G296-14 AF395037.1 91 Cab (A, I) Sphingomonas sp. KT-1 AB022601.1 94 Cab (A, I) Sporolactobacillus inulinus D16284.1 99 Sem (A, I) Yersinia sp. KM16 AJ011333.1 97 Sz (A, III) a, Sz, Shiraz, Cab, Cabernet Sauvignon; Mer, Merlot; Cha, Chardonnay; Sem, Semillon; Sab, Sauvignon Blanc b, A1, Vineyard A, 2001-2002; A, vineyard A, 2002-2003; B, vineyard B, 2002-2003; C, vineyard C, 2002-2003; D, vineyard D, 2002-2003; E, vineyard E, 2002-2003; F, vineyard F, 2002-2003 c, Maturity stages are referred to in Table 3.2

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APPENDIX 2 Ribosomal DNA sequence identification of 16S rDNA DGGE bands obtained from wine grapes

Name Accession Sequence Samplesa Number similarity (Vineyard b, Maturity stage c) (%) Acetobacter diazotrophicus clone AY230815.1 94 Cab(A1, V), Cha (A1, V), Sem (A1, V) Acinetobacter johnsonii AY167841.1 90 Sz (C, III) Acinetobacter radioresistens Z93445.1 88 Sz (C, IV) Acinetobacter sp. AB192395.1 92 Mer (E, III) Agrobacterium tumefaciens AY435491.1 93 Sz (A1, IV) Bacillus benzoevorans AY167801.1 93 Sz (C, V) Bacillus benzoevorans AY167808.1 98 Sz (D, I-IV), Cha (D, I-V), Cab (E, I), Cab (E, II), Cab (E, III), Cab (E, IV), Cab (E, V), Sz (F, I-V), Cha (F, I-V) Bacillus benzoevorans D78311.1 91 Sz (B, II), Cha (B, II), Cha (B, III), Cha (A, V) Bacillus circulans AY647299.1 91 Cab (E, IV) Bacillus drentensis AY466403.1 99 Sz (C, IV) Bacillus maroccanus X60626.1 93 Mer (E, II) Bacillus megaterium AJ616728.1 97 Mer (E, III) Bacillus sp. AY269870.1 93 Sz (C, IV) Bacillus sphaericus Y150451.1 94 Sz (C, IV) Bacterial species 16S rRNA gene Z73441.1 90 Sem (A, II) Bacterium PSD-1-6 AY822573.1 96 Cab (A1, V) Bradyrhizobium sp. AF510604.1 97 Sem (C, II) Brevibacterium equis AJ251780.1 93 Sz (A1, V) Curtobacterium flaccumfaciens pv. AF273208.1 92 Sz (A1, V), Cab (A1, V), Cha (A1, beticola III), Cha (A1, V), Sem (A1, V), Sz (D, IV) Curtobacterium flaccumfaciens strain AF526909.1 93 Sz (A1, V), Sem (A1, V) 48v2 Curtobacterium flaccumfaciens strain AY67859.1 92 Sz (A1, V), Sem (A1, V) SAFR-001 Deinococcus radiodurans M21413.1 91 Sem (A, IV) Endosymbiont of Brevipalpus lewisi AB116515.1 90 Sz (B, III-V), Cha (B, III-V) Erwinia herbicola AB004757.1 89 Sz (C, III) Erwinia sp. AY690718.1 95 Sem (C, III) Flavobacterium sp. EP239 AF493659.1 96 Sem (A, III) Frankiaceae 16S ribosomal RNA X92365.1 93 Mer (E, IV) Friedmanniella sp AF409013.1 94 Sem (C, II) Geobacillus tepidamans AY563003.1 98 Sz (C, IV), Sz (C, V) Gluconobacter cerinus X80775.1 87 Cab (A1, V), Cha (A1, V), Sem (A1, V) Gluconobacter oxydans AB178431.1 96 Sz (A1, V), Cab (A1, V), Cha (A1, V), Sem (A1, V) Lactobacillus kunkeei Y11374.1 96 Cab (A1, V), Cha (A1, V) Lactococcus lactis AY348313.1 89 Sz (F, III) Lactococcus lactis AY675242.1 98 Sz (B, V), Cha (B, V) Methylobacterium sp. AB027621.1 99 Sem (C, IV) Microbacterium imperiale AF526916.1 93 Sem (C, IV), Cha (F, III) Microbacterium imperiale strain 27v1b AF526919.1 91 Sz (A1, V) Microbacterium paraoxydans AJ491806.1 88 Cha (A, III) Microbacterium sp. AF323562.1 90 Sz (C, IV)

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Name Accession Sequence Samplesa Number similarity (Vineyard b, Maturity stage c) (%) Mycobacterium sp. AY215227.1 90 Cab (E, IV) Nostoc sp. AY742454.1 98 Mer (E, IV) Paenibacillus dendritiformis AY359885.1 87 Sem (A, III) Pantoea agglomerans AF157694.1 93 Sem (A1, V) Pantoea sp. AY297724.1 97 Sem (C, II) Pseudomonas alcaligenes strain IH8-19 AY789554.1 92 Sz (A1, III) Pseudomonas putida AY456706.1 93 Sem (A1, V), Sem (C, IV) Pseudomonas putida U71007.1 97 Cha (A, III) Pseudomonas sp. AB079096 91 Cha (A1, III) Pseudomonas sp. AY267512.1 90 Sz (A, III) Pseudomonas sp. AF098465.1 98 Cha (A, III) Pseudomonas sp. AB079096.1 91 Cha (A, III) Pseudomonas sp. Fa20 AY131224.1 94 Cab (A1, V) Pseudomonas viridiflava AY604848.1 97 Sz (A, III), Cha (A, III), Sem (A, III), Sem (A, IV) Rhodocista sp. AR2107 AJ401217.1 92 Sz (A1, III) Serratia proteamaculans AJ279052.1 96 Sz (C, IV) Sphingomonas sp. AY167827.1 96 Cha (A1, III), Cha (A1, V) Sphingomonas sp. AY131296.1 97 Cab (A, II), Cab (A, III), Cab (A, V) Sphingomonas sp. SAFR-042 AY167827.2 96 Cab (A1, V), Sem (A1, V), Sem (C, V) Sphingomonas terrae D84531.1 100 Sem (C, III) Staphylococcus saprophyticus Z26902.1 98 Sem (C, III) Staphylococcus sp. AY635869.1 96 Sem (A, II) Staphylococcus sp. AF333342.1 96 Cha (D, III) Streptococcus thermophilus AY675258.1 98 Sem (C, V) Streptomyces sp. AF131504.1 89 Sem (C, IV) Streptomyces sp. AF128874.1 87 Cab (E, IV) Streptosporangium sp. AB124264.1 95 Sem (A, III) Uncultured actinomycete AJ555226.1 97 Cab (A, IV) Uncultured bacterium gene AB062145.1 95 Sem (A, II), Sab (E, III) Uncultured bacterium gene AB062145.1 98 Sem (C, III), Sem (C, V) Uncultured bacterium OIm26 AF018640.1 94 Cha (B, III) Uncultured earthworm intestine AY154503.1 88 Sem (A, III) bacterium clone Uncultured Methylobacterium sp. clone AY494631.1 98 Cha (B, III) Uncultured Ralstonia sp. clone AY734545.1 91-93 Sz (A1, III), Cab (A, IV), Sz (C, CDBL_C3 III), Sz (C, IV), Sem (C, III), Sem (C, V) Uncultured rape rhizosphere bacterium AJ295498.1 92 Mer (E, IV) Streptomyces scabiei AY207599.1 87 Sab (E, IV) Planococcus psychrotoleratus AY771711.1 87 Sab (E, IV) Uncultured earthworm cast bacterium AY154592.1 90 Sab (E, IV) clone Pseudoalteromonas spongia AY769918.1 91 Sab (E, V) Methylobacterium sp. AY436808.1 97 Sab (E, V) Tsukamurella tyrosinosolvens AY254699.1 97 Sab (E, V) Mycobacterium tusciae AF547978.1 95 Sab (E, III) uncultured Actinobacteridae bacterium AJ555207.1 92 Sab (E, V) a, Sz, Shiraz, Cab, Cabernet Sauvignon; Mer, Merlot; Cha, Chardonnay; Sem, Semillon; Sab, Sauvignon Blanc; b, A1, Vineyard A, 2001-2002; A, vineyard A, 2002-2003; B, vineyard B, 2002-2003; C, vineyard C, 2002-2003; D, vineyard D, 2002-2003; E, vineyard E, 2002-2003; F, vineyard F, 2002-2003; c, Maturity stages are referred to in Table 3.2

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APPENDIX 3 G 3.1 Schedule of pesticide application for vineyards from which grapes were sampled for bacterial species analysis (2001-2002 season)

Date of Vineyard Grape cultivar Pesticide applications* sampling Bravo 720, Bayfidan 250EC, 16/10/01 Sz, Cab, Ch, Sem Delfin WG 15/11/01 Sz, Cab, Ch, Sem Oxydule, Thiovit DF, Delfin WG Kocide Blue DF, Delfin WG, A 03/01/02 Sz, Cab, Ch, Sem Fortress 500 Dithane DF, Rovral Liquid, 30/01/02 Sz, Cab, Ch, Sem Delfin WG 10/02/02 Sz, Cab, Ch, Sem No applications * Applied approximately 1-14 days prior to sampling Sz, Shiraz; Cab, Cabernet Sauvignon; Ch, Chardonnay; Sem, Semillon Data supplied by viticulturists from various vineyards

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3.2 Schedule of pesticide application for vineyards from which grapes were sampled for bacterial species analysis (2002-2003 season)

Date of Vineyard Grape cultivar Pesticide applications* sampling Bravo 720, Bayfidan 250EC, 15/10/02 Sz, Cab, Ch, Sem Delfin WG 27/11/02 Sz, Cab, Ch, Sem Oxydule, Thiovit DF, Delfin WG Kocide Blue DF, Delfin WG, A 03/01/03 Sz, Cab, Ch, Sem Fortress 500 Dithane DF, Rovral Liquid, 24/01/03 Sz, Cab, Ch, Sem Delfin WG 31/01/03 Sz, Cab, Ch, Sem No applications Oxydule, Thiovit Jet MG, Dipel 15/10/02 Sz, Sem Forte Oxydule, Thiovit Jet MG, Dipel 27/11/02 Sz, Sem C Forte 03/01/03 Sz, Sem Oxydule, Dipel Forte 24/01/03 Sz, Sem Kocide Blue DF, Rovral Liquid 31/01/03 Sz, Sem No applications Topass, Polyram, Success, Dipel 25/10/02 Sz, Ch Forte Thiovit Jet, Kocide Blue, Dipel 05/12/02 Sz, Ch Forte D 09/01/03 Sz, Ch Rovral Liquid 02/02/03 Sz, Ch No applications 06/02/03 Ch No applications 20/02/03 Sz No applications Scala, Topass, Bravo, Polyram, 25/10/02 Cab, Mer, Sab Dipel Forte 05/12/02 Cab, Mer, Sab Spin, Dipel Forte E 09/01/03 Cab, Mer, Sab No applications 02/02/03 Cab, Mer, Sab No applications 06/02/03 Sab No applications 20/02/03 Cab, Mer No applications 29/10/02 Sz, Ch Sulfur, Captan, BT 28/11/02 Sz, Ch Flint, Spindo, BT 06/01/03 Sz, Ch Captan, Sulfur F 30/10/03 Sz, Ch No applications 04/02/03 Ch No applications 19/02/03 Sz No applications * Applied approximately 1-14 days prior to sampling Sz, Shiraz; Cab, Cabernet Sauvignon; Ch, Chardonnay; Sem, Semillon Data supplied by viticulturists from various vineyards

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APPENDIX 4 G Commercial name, active ingredient and application purpose for various pesticides used during wine grape cultivation

Commercial name Active ingredient Pest control Thiovit Jet Sulfur Mites and powdery mildew Delan 700 WG Dithianol Downy mildew and black spot Kocide Blue Copper hydroxide Frost Topas Penconazole Downy mildew and rusts Polyram DF Metiram Downy mildew and black spot Bravo Chlorothaloni Botrytis and downy molded Scala Pyrimethanil Botrytis Spin-Flo Benzothiadiazole Botrytis Oxydul Copper oxychloride Downy mild dew Rovral Liquid Iprodione Botrytis Spin Phenmedipham Control of weeds such as kochia, lambsquaters, mustard, green foxtail Fortress 500 Chlorethoxyphos Botrytis Dithane DF Mancozeb Downy mildew Rovral Iprodione Botrytis Proclaim Emamectin benzoate Moth in grapes Flint Trifloxystrobin Pest control Dipel Forte Bacillus thuringiensis Control certain leaf-eating caterpillars; gypsy subsp. kurstaki strain moth, spruce budworm, jack pine bud worm, EG2371 hemlock looper, tussock moth, tent caterpillar, Success Bacillus thuringiensis pine processionary moth, and others BT Bacillus thuringiensis Delfin WG Bacillus thuringiensis Data in this table was collected from http://www.awri.com.au/agrochemicals/right_chemical/recommendations.asp accessed on 15 August 2005 on the page of Agrochemicals Navigation of the Australian Wine Research Institute. G

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APPENDIX 5

5.1 Isolation of non-lactic acid bacteria by enrichment culture from Cabernet Sauvignon and Merlot grapes

Enrichment condition Grape Physical Vineyarda Autoenrichment MRS MRS + EtOH MRST (pH 5.5) MRST (pH 3.5) variety statusb 5 d 10 d 5 d 10 d 5 d 10 d 5 d 10 d 5 d 10 d Non-LAB Non-LAB Non-LAB Non-LAB Non-LAB Non-LAB UD - - - - (1.2u104)c (1.6u107) (6.2u105) (5.8u105) (5.0u103) (8.2u106) A Non-LAB Non-LAB Non-LAB Non-LAB Non-LAB Non-LAB D - - - - (7.2u106) (9.0u106) (4.0u107) (4.0u105) (1.0u107) (5.8u106) Bacillus sp. Bacillus sp. UD ------Cabernet (5.0u104) (2.9u106) D Sauvignon Non-LAB Non-LAB D ------(6.0u101) (2.4u106) Sp. inulinus Sp. inulinus Sp .inulinus Sp. inulinus Sp. inulinus Sp. inulinus UD - - - - (4.0u107) (9.0u105) (2.2u105) (2.5u104) (7.5u105) (4.2u106) F Sp. inulinus Sp. inulinus Sp. inulinus Sp. inulinus Sp. inulinus D - - - - - (1.2u107) (2.4u104) (1.4u106) (2.4u104) (1.4u106) UD ------Merlot D Non-LAB Non-LAB Non-LAB Non-LAB Non-LAB Non-LAB Non-LAB Non-LAB D (4.0u106) (9.0u106) (3.0u107) (7.0u106) (3.5u106) (4.9u107) (1.2u106) (1.8u106) a Vineyard location; A, Hunter Valley; D, Mudgee; F, Griffith; b UD, undamaged grape berries; D, damaged grape berries;c population (CFU/mL) in enrichment culture. Sp., Sporolactobacillus

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5.2 Isolation of non-lactic acid bacteria by enrichment culture from Pinot Noir and Tyrian grapes

Enrichment condition Grape Physical Vineyarda Autoenrichment MRS MRS + EtOH MRST (pH 5.5) MRST (pH 3.5) variety statusb 5 d 10 d 5 d 10 d 5 d 10 d 5 d 10 d 5 d 10 d Non-LAB Non-LAB Non-LAB Non-LAB Non-LAB Non-LAB Non-LAB Non-LAB Non-LAB Non-LAB UD (4.0u106)c (5.5u106) (8.7u106) (5.6u106) (1.2u103) (3.0u102) (1.0u107) (8.5u107) (1.0u106) (3.3u106) Pinot Non-LAB D Noir Non-LAB Non-LAB (2.1u107) Non-LAB Non-LAB Non-LAB Non-LAB Non-LAB D - - (1.7u106) (2.4u106) Bacillus sp. (3.1u107) (5.4u106) (1.1u107) (4.4u106) (4.4u106) (3.0u105) UD Sp. inulinus Sp. inulinus Sp. inulinus Sp. inulinus ------(1.9u107) (1.0u103) (2.0u101) (8.9u103) Tyrian F D Sp. inulinus Sp. inulinus Sp. inulinus Sp. inulinus Sp. inulinus - - - - - (1.8u106) (2.0u102) (8.0u102) (9.5u104) (4.5u105) a Vineyard location; D, Mudgee; F, Griffith; b UD, undamaged grape berries; D, damaged grape berries; c population (CFU/mL) in enrichment culture. Sp., Sporolactobacillus

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5.3 Isolation of non-lactic acid bacteria by enrichment culture from Shiraz grapes

Enrichment condition Physical Vineyarda Autoenrichment MRS MRS + EtOH MRST (pH 5.5) MRST (pH 3.5) statusb 5 d 10 d 5 d 10 d 5 d 10 d 5 d 10 d 5 d 10 d Non-LAB Non-LAB Non-LAB Non-LAB Non-LAB (1.7 106) UD - - - - - u (1.1u107) (1.1u107) (1.3u107) (1.0u104) Non-LAB (2.3 106) A u Non-LAB Non-LAB Non-LAB Non-LAB Non-LAB Non-LAB Non-LAB Non-LAB (5.0 105) (4.3 105) D - - u u (4.2u105) (6.3u106) (1.7u107) (8.1u105) (2.2u106) (1.3u107) Non-LAB Non-LAB (5.0u105) (4.0u105) A. siamensis A. siamensis Non-LAB A. siamensis A. siamensis A. siamensis UD - - - - (4.0u105) (4.0u105) (3.6u103) (1.0u105) (1.6u105) (3.1u106) B A. siamensis A. siamensis D ------(1.2u106) (1.5u106) Non-LAB Non-LAB UD ------C (2.1u107) (9.0u105) D ------S. capitis S. capitis UD ------(3.7u105) (5.4u105) D Non-LAB Non-LAB Non-LAB Non-LAB Non-LAB Non-LAB Non-LAB Non-LAB Non-LAB Non-LAB D (6.3u106) (1.5u105) (5.4u107) (4.9u106) (1.8u106) (1.1u108) (1.6u107) (1.7u108) (1.6u106) (2.2u105) UD ------E D ------S. capitis Sp. inulinus Sp. inulinus Sp. inulinus Sp. inulinus Sp. inulinus UD - - - - (4.3u105) (4.1u105) (8.2u102) (1.5u107) (2.1u102) (8.0u104) F A. siamensis A. siamensis D ------(1.8u104) (5.9u106) a Vineyard location; A, B, C, Hunter Valley; D, E, Mudgee; F, Griffith; b UD, undamaged grape berries; D, damaged grape berries; c population (CFU/mL) in enrichment culture. A., Asaia; S. Staphylococcus; Sp., Sporolactobacillus G G 210

5.4 Isolation of non-lactic acid bacteria by enrichment culture from Chardonnay grapes

Enrichment condition Physical Vineyarda Autoenrichment MRS MRS + EtOH MRST (pH 5.5) MRST (pH 3.5) statusb 5 d 10 d 5 d 10 d 5 d 10 d 5 d 10 d 5 d 10 d UD ------A Non-LAB Non-LAB Non-LAB Non-LAB Non-LAB Non-LAB D - - - - (1.0u106) (1.0u107) (1.0u106) (4.8u106) (8.3u105) (1.2u107) Non-LAB Non-LAB Non-LAB Non-LAB UD ------(1.0u106) (1.7u107) (1.7u107) (1.7u107) C Non-LAB Non-LAB Non-LAB Non-LAB Non-LAB G.. oxydans G.. oxydans D - - - (2.9u106) (1.7u103) (4.0u106) (1.0u108) (2.0u106) (1.0u107) (7.0u107) UD ------D D ------S. hominis S. hominis UD ------E (5.0u105) (2.8u107) D ------Sp. inulinus Sp. inulinus (2.4u105) (4.4u106) UD ------Staphylococcus Staphylococcus F sp. (2.0u105) sp. (1.2u106) Non-LAB Non-LAB D ------(6.4u105) (3.4u106) a Vineyard location; A, C, Hunter Valley; D, E, Mudgee; F, Griffith; b UD, undamaged grape berries; D, damaged grape berries; c population (CFU/mL) in enrichment culture. S., Staphylococcus; Sp., Sporolactobacillus

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5.5 Isolation of lactic acid bacteria by enrichment culture from Semillon and Sauvignon Blanc grapes

Enrichment condition Grape Physical Vineyarda Autoenrichment MRS MRS + EtOH MRST (pH 5.5) MRST (pH 3.5) variety statusb 5 d 10 d 5 d 10 d 5 d 10 d 5 d 10 d 5 d 10 d A. siamensis A. siamensis A. siamensis A. siamensis A. siamensis A. siamensis A. siamensis UD - - - (2.9u103) (5.4u106) (3.2u106) (1.0u105) (1.0u107) (1.0u105) (2.0u102) A Non-LAB Non-LAB Non-LAB Non-LAB Non-LAB Non-LAB Non-LAB D - - - (8.2u106) (5.0u106) (2.6u106) (1.0u105) (1.0u107) (4.5u105) (3.6u106) UD ------Semillon B Non-LAB Non-LAB D ------(1.2u106) (3.6u106) Staphylococcus Staphylococcus Staphylococcus Staphylococcus UD ------F sp. (3.1u106) sp. (1.0u104) sp. (4.8u104) sp. (7.5u105) D ------Non-LAB Non-LAB Non-LAB Non-LAB Non-LAB Non-LAB Non-LAB Non-LAB UD - - (2.0u106) (3.3u106) (6.1u106) (4.9u107) (5.0u106) (2.3u107) (4.7u104) (1.5u106) Sauvignon D Non-LAB Non-LAB Blanc D ------(3.0u106) (7.0u106) F D ------a Vineyard location; A, B, Hunter Valley; D, Mudgee region; F, Griffith region; b UD, undamaged grape berries; D, damaged grape berries; c population (CFU/mL) in enrichment culture; A., Asaia.

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