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thesis entitled

MANAGEMENT OF MICROBIAL DECAY OF FRESH AND
PEELED CHESTNUTS IN MICHIGAN

presented by

IRWIN R. DONlS-GONZALEZ

has been accepted towards fulﬁllment
of the requirements for the

MASTER OF degree in PLANT PATHOLOGY
SCIENCE

 

 

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MANAGEMENT OF MICROBIAL DECAY OF FRESH AND PEELED
CHESTNUTS IN MICHIGAN

By

Inivin R. Donis-Gonzélez

A THESIS
Submitted to
Michigan State University
In partial fulﬁllment of the requirements
For the degree of

MASTER OF SCIENCE

Plant Pathology

2008

Copyright by
Irwin R. Donis—Gonzalez
2008

ABSTRACT

MANAGEMENT OF MICROBIAL DECAY OF FRESH AND PEELED
CHESTNUTS IN MICHIGAN

By
Irwin R. Donis-Gonzalez

Edible chestnuts (Castanea spp.) represent a worldwide growing
commodity. Worldwide including Michigan, postharvest losses due to microbial
activity, including several ﬁlamentous fungi are problematic for the industry. To
determine the organisms involved with shell mold and kernel decay, a survey for
microorganisms associated with fresh chestnut was performed. Eleven species
of molds were found to negatively impact fresh chestnuts; two species had never
been found on chestnut prior to this study. Microbial populations were dependent
on harvest methods and the farms from which the chestnuts were collected. A
survey of peeling equipment to determine the source of contaminants on peeled
chestnuts showed that the skin separator and the sorting belt were sources of
contamination by two bacterial species and one species of yeast. To reduce
microbial growth on fresh and peeled chestnut, several sanitizers were
evaluated. Hydrogen peroxide and triﬂoxystrobin signiﬁcantly reduced shell mold
severity and kernel incidence on fresh chestnuts. X-ray irradiation, hydrogen
peroxide and hot water immersion signiﬁcantly reduced spoilage on peeled
chestnuts. Information from this study adds levels of protection against
postharvest chestnut mold, decay and spoilage when combined with other good

manufacturing and agricultural practices.

To my parents, sister, brother and Yvonne, who can never know how much their
love, encouragement and moral support has meant to me through the
development of this thesis. With lots of love this is for you...

ACKNOWLEDGMENTS

I owe a debt of gratitude to many people for their support and contributions
to this thesis.

First, I would like to thank my advisor, Professor Dennis W. Fulbright, who
developed the initial idea for the project and contributed excellent guidance and
moral support throughout. Professor Daniel E. Guyer and Professor Elliot Ryser
who also made innumerable useful suggestions, experimental design,
development and editorial comments.

I would also like to thank Professor Ray Hammerschmidt, Dr. Zhinong
Yan, Dr. Sandhyup Jeong, Dr. Jianjun Hao, Dr. Phillip Wharton, Professor
Annemiek Schilder and Professor Francis Trail; highly known scientist from
diverse departments at Michigan Sate University, for their technical and
knowledge support, during the experimental design and development of diverse
studies.

I am particularly grateful to all who provided ﬁnancial support during my
studies at Michigan State. The US. Department of State through the Fulbright
Scholarship Program, Michigan State University and Greeen Grant Program and
The Michigan Chestnut Grower Incorporation, who provided much support during
my career and all stages of the study.

Finally, I would like to express my deepest appreciation to everybody who
participated in the study and also offered countless hours of moral support,

including Sara Stadt, Mario Mandujano, Carmen Medina, Virginia Rinkel, Marion

Heller, La Fever Family and fellow students. Without their assistance and help
provided, I could obviously never have performed this study.

Perhaps the largest share of my appreciation goes to my family and
Yvonne. Without their loving faith, this study wont not have been possible, nor

worth doing.

TABLE OF CONTENTS

LIST OF TABLES - viii

LIST OF FIGURES - ix

CHAPTER 1. LITERATURE REVIEW

 

 

 

 

 

 

 

Introduction 1
Chestnut fruit 6
World chestnut industry 9
Postharvest organisms in fresh and peeled chestnuts 13
Sanitizers, postharvest and post processing treatments 20
Objectives of this study 29
REFERENCES 31

CHAPTER 2. SHELL MOLD AND KERNEL DECAY OF FRESH CHESTNUTS IN
MICHIGAN

 

 

 

 

 

 

 

 

 

 

 

 

 

 

Abstract 44
Introduction 45
Collection and storage of chestnut samples 47
Microbial populations - First and third study 49
She" 49
Kernel 49
Microbial mold severity and kernel decay incidence analysis - Second study .......... 50
Identiﬁcation of microorganisms - Third study 50
Statistical analysis 52
Results 53
Shell mold symptoms 53
Kernel decay symptoms 59
Inﬂuence of chestnut harvesting method on microbial populations 62
Farm inﬂuence on microbial populations on she" and in the kernel of chestnuts...66
Farm inﬂuence on microbial quality of chestnuts 69
Discussion 71
REFERENCES 75

 

CHAPTER 3. EFFICACY OF POSTHARVEST TREATMENTS FOR REDUCTION OF
MOLD AND DECAY IN FRESH CHESTNUTS

 

 

 

 

 

 

Abstract 80
Introduction 81
Material and Methods 83
Samples 83
Treatments and storage 84
Microbial mold severity, kernel incidence and weight loss analysis 87
Statistical analysis 88

 

vi

Results 88
Discussion 90

REFERENCES ................... 95

 

CHAPTER 4. MICROBIAL CONTAMINATION IN PEELED CHESTNUTS AND THE
EFFICACY OF POST-PROCESSING TREATMENTS FOR MICROBIAL SPOILAGE

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

MANAGEMENT
Abstract 102
Introduction 103
Material and Methods 106
Chestnut samples 106
Environmental and water samples 106
Determination of microbial populations in fresh chestnut, peeled chestnut and
environmental samples 108
Identiﬁcation of microorganisms 110
Determining symptoms associated with microorganisms isolated from spoiled peeled
chestnuts 110
Statistical analysis 115
Results 116
Microorganisms associated with spoiled peeled chestnuts 116
Microbial population 118
Post-processing treatments 122
Discussion 125
REFERENCES 131
APPENDICES ................. 138
APPENDIX A. Mycotoxins in fresh Michigan chestnuts 139
APPENDIX B. Commercial-level peeling system (Boema; Neive, Italy) ........ 142
APPENDIX C. Chestnut fruit morphology 144
APPENDIX D. World chestnut production ..... 146
APPENDIX E. Inhibition properties of pellicle, hilum, shell and kernel against
Cladosporium cucumerinum 148

 

vii

LIST OF TABLES

Table 1. Advantages, disadvantages and uses of some available sanitizing
agents, salts, chemicals, bio-control agents, hot water and irradiation for
postharvest and postprocessing treatment of fresh fruits, vegetables and foods

....................................................................................................................... 23
Table 2. Microorganisms isolated from stored chestnut shell and kernel during
the 2006 and 2007 growing seasons in Michigan .......................................... 55
Table 3. Postharvest treatments used in fresh chestnuts .............................. 84

Table 4. Effect of fresh chestnut treatment application on shell mold, kernel
decay and weigh loss stored for 60 days in control environment ................... 89

Table 5. Post-processing treatments used in artiﬁcially inoculated peeled
chestnuts ...................................................................................................... 1 15

viii

LIST OF FIGURES

Figure 1. Chestnuts showing shell mold ........................................................... 59
Figure 2. Chestnuts showing kernel decay ........................................................ 62

Figure 3. Mean mold counts (log cfu g") on shells and in kernels of fresh
chestnuts during 120 days of storage at 4° C. .................................................... 64

Figure 4. Mean MAB counts (log cfu g") on shells and in kernels of fresh
chestnuts during 120 days of storage at 4° C. .................................................... 65

Figure 5. Mean yeast counts (log cfu g") on shells and in kernels of fresh
chestnuts during 120 days of storage at 4° C. .................................................... 66

Figure 6. Total MAB, molds and yeast counts (log cfu g") from the shells of fresh
chestnuts after harvest from seven Michigan farms. .......................................... 68

Figure 7. Total MAB, molds and yeast counts (log cfu g") from the kernels of
fresh chestnuts after harvest from seven Michigan farms. ................................. 69

Figure 8. Shell mold severity and kernel decay incidence from seven Michigan

farms at receiving day (Day 0). ........................................................................... 70
Figure 9. Shell mold severity and kernel decay incidence from seven Michigan

farms, during 60 day storage at 4°C. .................................................................. 71
Figure 10. Treatment of fresh chestnuts before storage .................................... 87
Figure 11. Sample collection points for fresh and peeled chestnuts. ............... 107

Figure 12. Schematic diagram of a brulage chestnut peeling line (Boema; Neive,
Italy), describing chestnuts, environmental and water sampling points ............ 108

Figure 13. Peeled chestnuts, 10 days after inoculation and stored at 4 °C. 117

Figure 14. Total MAB, molds and yeast counts (log cfu g") on peeled chestnuts
before and during peeling process (Brulage peeler - Boema; Neive, Italy). ..... 119

Figure 15. Total MAB, molds and yeast counts (log cfu/cmz) on environmental
samples during peeling process (Brulage peeler - Boema; Neive, Italy). ......... 120

Figure 16. Total MAB, molds and yeast counts (log cfu g“) in water samples
during peeling process (Brulage peeler - Boema; Neive, Italy). ........................ 122

Figure 17. Mean MAB and yeast counts (log cfu g") from peeled chestnuts
treated with different sanitizers during 18 days of storage at 4° C .................... 123

Figure 18. Mean MAB and yeast counts (log cfu g") from peeled chestnuts
treated with different X-ray irradiation doses during 18 days of storage at 4° C..

.......................................................................................................................... 125
Figure 19. Presence of mycotoxins in two cultivars of fresh Michigan chestnut:40
Figure 20. Schematic diagram of a brulage chestnut peeling line (Boema; Neive,

Italy) .................................................................................................................. 143
Figure 21. Chestnut fruit morphology. .............................................................. 145
Figure 22. World chestnut (Castanea spp.) production in 2001 ...................... 147

Figure 23. Inhibition properties of pellicle, hilum and shell against Cladosporium
cucumerinum. ................................................................................................... 149

CHAPTER 1

LITERATURE REVIEW

Introduction

Chestnut (Castanea spp.) is one of the most popular nut-bearing trees in
the Mediterranean region, as well as many countries of Asia including China,
Japan and Korea (Ridley, 1999). Chestnuts grown in Europe and Asia account
for more than 70% of the worldwide production with chestnut production currently
increasing in Australia, New Zealand and Chile (Fulbright and Mandujano, 2000;
Grau and France, 1999; Klinac et al., 1999; Ridley, 1999; Mencarelli, 2001). In
the United States chestnuts are rare; people are more likely to be familiar with the
unrelated and poisonous horse chestnut (Aesculus spp.) (Fulbright and
Mandujano, 2000) than with any of the edible, sweet chestnut species (C. dentata
Borkh., C. mollissima BI., C crenata Sieb and Zucc., C. sativa Mill., C. seguinii
Dode, C. pumila Mill, C. henryi Rehd. and VIﬁls.) (Anagnostakis et al., 1998;
Fulbright and Mandujano, 2000; Miller, 2003). This is partially due to chestnut
blight (Cryphonectn’a parasitica Murril and Barr), which virtually eliminated the
once-widespread American chestnut (C. dentata) during the ﬁrst half of the
twentieth century (Merkel, 1905; Metcalf, 1912; Gravatt and Marshall, 1926).
Despite advances in chestnut blight research, chestnut blight still remains one of
the major limitations to growing American chestnuts (Fulbright and Mandujano,
2000; MacDonald and Double, 2006; Anagnostakis et al., 1998). Nevertheless,

commercial orchards have been established in different locations in the United

States, and have been increasing over the years (Fulbright and Mandujano,
2000). In the past, especially in Europe and in China, chestnut consumption was
predominant in rural areas and was considered as a food for the poor. Currently,
due to their unique nutritional value, chestnuts are broadly consumed and used
as a cooking ingredient with health-related beneﬁts (De la Montana Miguelez et
al., 2004; Borges et al., 2008; Xu, 2005). Recently, the number of chestnut
plantations has increased in the United States, including Michigan, and both fresh
and peeled domestic chestnuts are becoming more common. Relatively recent
domestic availability of chestnuts has triggered an increase in consumption.
Vossen (2000) claimed that the United States has a great potential for chestnut
production and a small rise in domestic consumption could be worth up to $800
million annually. Similar to patterns in Europe, chestnuts consumed in the United
States, are most commonly prepared by roasting the fresh product (Harte et al.,
2003) but other diverse culinary applications are also available (CGI: Chestnut
Growers, Inc 2008; Kelley and Behe, 2002). In studies performed in Wisconsin
and California to evaluate consumer-preferences for chestnuts, between the
years 2000 and 2004, most of those interviewed had never tasted a chestnut, but
were interested in exploring them as a new food. Quality and nutritional value
were listed as the most important attributes inﬂuencing purchase and
consumption decisions (Gold et al., 2004; Vossen, 2000).

The quality of fresh chestnuts is limited due to microbial decay after
harvest and during storage. Organisms that cause postharvest decay of fresh
commodities are found worldwide and in most cases, it has been estimated that

losses due to postharvest decay could be greater than 25 percent of all harvested

foods. Under poor storage conditions, and if adequate precautions are not taken,
microbial growth is favored and losses caused by postharvest diseases may be
greater than the economic value of the non-decayed portion of the crop
(Narayanasamy, 2006). Chestnuts are not an exception. Destruction by various
ﬁlamentous fungi, commonly called molds, is an important and predominant
factorin worldwide economic and postharvest losses (Jerimini et al., 2006;
McCarter et al., 1980; Narayanasamy, 2006).

Postharvest decay of chestnuts reduces product quality and leads to
unmarketable chestnuts. Although a certain amount of affected product may be
sold in less demanding markets, this practice results in important economic and
potential market losses due to consumer rejection. A number of mold-s can cause
postharvest decay and disease problems in chestnuts. Among these, Penicillium
spp., Aspergillus spp., Fusan'um spp., Phomopsis castanea, Acrospeira mirabilis,
and Sclerotinia pseudotuberosa have been repeatedly identiﬁed in France, Italy,
Australia, Chile, United States and other countries (Ellis and Ellis, 1985; Jerimini,
et al., 2006; Montealegre, J. and Gonzalez, S. 1986; Paglietta and Bonous, 1979;
Ridé, M. and Gudin, C. 1960; Vettraino et al., 2005 Washington at al. 1997;
Wright, 1960). In 2007, more than 25 percent of the total Michigan crop was lost
due to postharvest decay, equivalent to approximately 11,500 pounds of fresh
chestnuts, which reﬂects a potential annual economic loss of approximately
$30,000 or more for the young Michigan start-up chestnut growers’ cooperative,
Chestnut Growers, lnc. (CGI).

Furthermore, some fungi are capable of secreting substances that are

potent, acute toxins or carcinogens to both animals and humans. These toxic

agents are called mycotoxins and their impact on domestic animals in terms of
decreased growth rate, abnormal reproduction and early death has long been
recognized (Adams and Moss, 2000; Bullerman and Bianchini, 2007; Cho et al.,
2008; Mateo and Jimenez, 2007; Schollenberger et al., 2008; Tanaka et al.,
2007; Varga, et al., 2007). The most important mycotoxins are aﬂatoxins,
deoxynivalenol (DON), zearalenone, fumonisin B1, T-2 toxin, and ochratoxin A.
These substances have been predominately isolated from grains such as wheat
and corn, but can also been found in chestnuts as well as in other products
including rice, grapes, beer, wine, vegetable oil, vegetables, peanuts, pecans and
processed foods (Adams and Moss, 2000; Bullerman and Bianchini, 2007; Cho et
al., 2008; Mateo et al., 2007; Lillard et al., 1970; Schollenberger et al., 2008;
Tanaka et al., 2007; Varga, et al., 2007). In a nationwide survey in Canada from
1998 to 1999, penitrem A, chaetoglobosin A and C, emodin and ochratoxin A
were found on imported chestnuts sold by grocery stores (Overy et al., 2003).

As an alternative to fresh chestnuts, peeled chestnuts can be sold as a
value-added product to extend the chestnut market beyond seasonal sales,
providing the opportunity to expand markets and their utilization. In Michigan,
peeled chestnuts can be obtained by processing fresh chestnuts with a
commercial peeling system obtained from Italy in 2001 (Boema; Neive, Italy)
(Appendix B). After peeling, the chestnuts are vacuum-packed and stored frozen
(Guyer et al., 2003). Other mechanical methods, including air-impingement de-
shelling are also available but are not used in Michigan (Gao et al., 2008). Peeled
frozen chestnuts, often develop a sticky, yellow ooze over the surface of the nut,

within twelve days after thawing that affects quality and acceptability (B. Harte

4

and D.W. Fulbright, personal communication). Similar problems have not been
previously reported in peeled chestnuts, but are common for processed fruits,
vegetables, juices, ready-to-use salads and meats. In general, the growth of
spoilage microbes, including several bacteria and yeasts, is usually accompanied
by the accumulation of metabolites, such as ethanol, lactic acid, and ethyl
acetate. These organoleptic changes due to microbial activity are associated with
the enzymatic oxidation of various compounds leaking from the injured tissues.
Spoilage is detectable by sensory and microbiological methods. When the
microbial population attains a speciﬁc level, spoilage is dependent on the species
in question and the ingredient assayed. In extreme cases, the refrigerated (4 °C)
shelf life of several commercial products may not exceed ﬁve days (Brightwell et
al., 2007; Guerzoni et al., 1996; James et al., 2005; N9, 2007; Ragaert et al.,
2006; Tournas, 2005).

All these issues regarding postharvest mold, decay, post-processing
spoilage of peeled chestnuts and safety concerns, reﬂect the need for more
effective microbial reduction strategies after harvest and processing. In response
to these needs, one speciﬁc goal of this research was to determine the potential
organisms that interact with both fresh and peeled chestnuts. Elucidating these
organisms will help assess the efﬁcacy of various physical, chemical, and
biological treatments for reducing the microbiological populations ensuring the

quality, safety and prolonged shelf-life of fresh and peeled chestnuts.

Chestnut fruit

Many true nuts, including chestnuts, are wrapped in a papery or spiny
husk called the involucre, more commonly known as a bur (Appendix C). This
structure is usually mistaken as the fruit, but it is actually a whorl of modiﬁed
leaves around the ﬂower or ﬂowers. Inside the involucre, depending on the
species or cultivar, one or several chestnuts can be found (usually three). In
chestnuts, the ovule or ovules that develop into a seed or several seeds are
called kernels, structures that are formed inside an ovary, which will swell and
give rise to what is known as the shell, but botanically is a fruit called an achene
(pericarp). This structure has an external brown wall with a woody and shinny
appearance. Two large cotyledons, forming the kernel, surround the embryo,
which contains the radicle, hypocotyls and epicotyls. Between the shell and the
kernel, is a thin brown papery-like structure, commonly called a pellicle, which is
botanically known as a seed coat, testa or episperrn (Mencarelli, 2001; Miller G.,
2003). Usually there is only one kernel per shell, but in certain cases there may
be two or more (commonly called a double embryo). In different regions of
Europe, including Italy and Spain, the term “marron” denotes a single kernel nut,
while the term “chataigne”, “castano” or “castafia” denotes double or even more
kernels per shell. Therefore, from a botanical point of view, a chestnut is a fruit
(achene) containing one or more seeds (kernel or edible part) with creamy,
yellow-colored cotyledons that are covered by a membrane called the pellicle

(episperm) (Mencarelli, 2001; Miller G., 2003).

When chestnuts start to ripen in late summer or autumn, the bur changes
color from light green to yellow-brown and releases the chestnuts. Sometimes the
bur opens on the tree, releasing the chestnuts, but the bur can also drop and
open on the ground (Anagnostakis et al., 1998; Mandujano et al., 1998;
Mencarelli, 2001; Miller, 2003; VWIis et al., 2007).

Unlike other nuts like almonds, hazelnuts and walnuts, the chestnut kernel
is starch-based (40 - 90 percent‘), slightly hard, containing high ﬁber (14 - 19
percent), protein (6 - 10 percent) and low lipids (0.4 - 10 percent), which are 90
percent unsaturated fatty acids. Chestnuts are also rich in sugars mainly sucrose,
glucose and fructose (40 -60 percent) and its water content is relatively high (>50
percent) (Anagnostakis and Devin, 1999; Fulbright, 2003). Furthermore,
chestnuts are a source of vitamin A, calcium, iron, ﬁber with and antioxidants
(Biomhoff et al., 2006; Gao et al., 2008). Biomhoff et al. (2006) and Anagnostakis
and Devin (1999) reported that all these characteristics are highly beneﬁcial for
human health and are needed for proper nutrition and protection of animal cells,
ranking chestnuts as a healthy food for consumers.

After harvest, respiration of most seeds is usually characterized by a low
and constant rate without any peak, reﬂecting a non-climacteric respiration
pattern. The same relative pattern can be found in chestnuts, but when compared
to other seeds and to most other nuts, chestnuts have a higher respiration rate.
Subsequently, due to their high respiration rate, water loss and starch conversion

to sugars is high (Kader, 2002; Willis et al., 2007). Based on several respiration

 

1 Nutrients in chestnuts expressed as the percentage of dry weight

rate studies (Harte et al., 2003), chestnuts were comparable to certain berries
(i.e., blueberries) (Perkins-Veasie, 2004) and iceberg lettuce (Smyth and
Cameron, 1998; Kader, 2002), and exhibited a lower respiration rate than out
broccoli (Talasila et al., 1994), peas, asparagus, sweet corn and mushrooms
(Kader, 2002).

Therefore, the most common changes that occur in chestnuts after harvest
are moisture loss, starch conversion to sugar, fungal decay and insect damage
(Wells, 1980). Only starch conversion to sugar (commonly called curing)
positively affects chestnut quality, enhancing the ﬂavor, sweetness and
acceptability of the product (Harte et al., 2003). The other three major factors
have a negative affect on quality and storage potential of the ﬁnal product (Harte
et al., 2003; Jerimini et al., 2006; Montealegre and Gonzalez, 1986; Paglietta and
Bonous, 1979; Wells, 1980). Because quality can only be maintained and not
improved after harvest (Kader, 2002; Willis et al., 2007), efforts have focused on
increasing the storability of chestnuts by reducing microbes and insects and
maintaining the quality at harvest. Vossen (2000), reported that during three
months storage, temperatures between -2 °C to 0.5 °C were recommended and
Dooley et al. (1980) and Jian et al. (2002) concluded that controlled atmosphere
(CA) plays an important role in
inhibiting fungal decay, increasing storage and reducing moisture loss. Currently,
storage practices by growers and cooperatives have been unable to efﬁciently

maintain chestnuts quality after harvest (Harte et al., 2003). This afﬁrms the

urgent need to develop better postharvest chestnut handling and storage

practices.

World chestnut industry

According to FAO in 2002, at least 25 countries produced chestnuts. VIﬁth
the exception of France, chestnut production has experienced continuous growth
(FAO, 2002; Vossen, 2000) during the past 10 years. Chestnut production
worldwide was estimated of over 500,000 tons (454,000 metric tons) during
2000-2001 and was distributed as follows: China, 23.7 percent; Korea, 23.3
percent; Italy, Turkey, and Japan, about 10-15 percent each; France, Spain, and
Greece, about 5 percent each; and the United States, Australia, New Zealand,
Chile, among others, less than 1 percent each (FAO 2002) (Appendix D). This
production pattern is mainly due to the increase in cultivation areas and changes
in consumer preferences towards healthier and more convenient foods (Gold et
al., 2004; Kader, 2002; Vossen, 2000).

China is the largest low-cost producer and exporter of chestnuts with an
estimated production of 117,000 tons (105,300 metric tons) and exports about a
third of their chestnuts to Japan. Most chestnuts are consumed fresh or roasted
with an unspeciﬁed amount used in Chinese cuisine to develop a broad variety of
dishes.

Korea, the second largest producer yields almost the same amount of
chestnuts as China per year (115,000 tons = 103,500 metric tons), approximately

half of which are exported to Japan and some to the United States.

Japan is the largest chestnut importer and is among the largest consumer
group, even though it is not the biggest producer. In Korea and Japan, local or
imported chestnuts are primarily stored under refrigerated conditions of 4 — 7 °C
and consumed fresh, boiled or as an ingredient in diverse dishes. Some are also
stored dry or peeled for further use.

VIﬁthin Europe, Italy is the largest chestnut producer and leads the world in
production of delicacy-processed products such as mamone glacé (preserved
chestnuts in sugared liquor). In Europe, the use of dry chestnuts and chestnut
ﬂour in cooking has recently declined, but the popularity of these products is
increasing elsewhere, especially in the United States. These value-added
products, such as processed, dried, peeled, and frozen chestnut have reached
more than US $3.00 per pound, prompting moves to expand the chestnut
industry (Vossen, 2000). Europe’s second largest producer is France, with up to
25,000 tons (23,000 metric tons) per year. Irrespective of the decline in
production over the past 10 years, due to other more valuable crops and
urbanization, France is one of the biggest importers of chestnuts in Europe,
mostly from Italy, Spain and Portugal.

Recently the United States, Australia, New Zealand, Chile and other
countries in the Southern Hemisphere have begun to produce chestnuts
and have established economically sustainable industries, mainly for export
(FAO, 2002; Vossen, 2000; Fulbright and Mandujano, 2000). The United States,
has at least of 2,500 acres of chestnut tree plantations. Of these, approximately

1,500 acres of young (less than 10 years) chestnut trees are distributed among

10

Michigan, Oregon, Washington and Ohio (Fulbright, 2008; Vossen, 2000). These
commercial plantations are primarily from the cultivar ‘Colossal’ (European-
Japanese hybrid = C. sativa x C. crenata). This cultivar has been broadly used,
because it produces large nuts (up to 30 9), high yields (> 55 kg/tree), and is
commercially sold at nurseries (Miller, 2003; Fulbright and Mandujano, 2000).
Other cultivars such as ‘Dunstan Hybrids’ (a patented seedling of a third
generation cross between American and Chinese chestnut hybrids = C. dentata x
C. mollissima), ‘Skookum’, ‘Layeroka’, ‘Myoka’, ‘Skioka’, ‘Eaton’ which have
apparent Chinese characteristics have also been planted (Miller, 2003; Fulbright
and Mandujano, 2000; Vossen, 2000).

In recent years chestnut average production has been increasing (Vossen,
2000). As a point of reference, in 2003, approximately 3,000 pounds of chestnuts
were harvested by the Chestnuts Grower Incorporation (CGI) in Michigan, while
in 2007 a total of 45,000 pounds were harvested (Blackwell, 2006; Fulbright D.
W., 2008). An important chestnut state is California where the oldest commercial
plantations are found. Vossen (2000) indicated that most of the early chestnuts
orchards established in California were brought by immigrants during the Gold
Rush and are mostly seeds from European chestnuts.

Michigan’s chestnut harvest starts in mid-September and proceeds
through the ﬁrst week of November but may be slightly later in northern or colder
locations. Product is primarily sold through CGI a producer owned and controlled
marketing cooperative with about 40 members. Most chestnuts are sold from
Thanksgiving through Christmas, via sales to specialty ethnic markets, retail

stores, food processors, restaurants, holiday festivals, farrners’ markets and

11

individual consumers. Fresh and frozen peeled chestnuts compose up to 60 and
30 percent, respectively of outlet sales. Most of the remaining chestnuts are sold
as ﬂour, breading and new dehydrated products such as chips and slices, and
puree (Blackwell, 2006; Fulbright D. W., 2008).

Since freshness and microbial quality are sales factors in chestnut
appearance, domestic production has a deﬁnite advantage over imported
chestnuts. Due to this advantage, storage conditions and postharvest
management strategies are both important in preventing the chestnuts from
molding and decaying. Regardless of quality and lack of freshness due to long-
term storage and transport of imported chestnuts, the United States annually
imports between 10 and 20 million pounds (4.5 to 9 million kg) of fresh European
and Chinese chestnuts, at a retail price of approximately 40 million dollars. In
order to the United States to expand its markets, replace imported chestnuts, and
fulﬁll its all local needs, more than 5,000 acres (2,000 ha) of production would be
required. Furthermore, if chestnuts are marketed efﬁciently and domestic
consumption increases by only half a pound (0.22 kg) per capita, the United
Sates would require over 50,000 acres (20,200 ha) of mature production to meet
their demand. Following this growth, the industry would be worth more that $300
million annually, but may be worth more than $800 million annually, if the
increase in consumption is even higher (Vossen, 2000).

Taken together, these suppositions indicate that the opportunity for
expanding chestnut markets is a feasible venture. Fulbright and Mandujano
(2000) indicated that establishing chestnut orchards in United States, including

Michigan, has not been an easy task, but if they are managed appropriately and

12

planning is done efﬁciently, it may be economically successful. Nevertheless,
regardless of the location, anyone considering investing in a chestnut orchard
should evaluate individual production costs, and earning potential, and is advised

to make comparisons with alternative investments.

Postharvest organisms in fresh and peeled chestnuts

Although Adams and Moss (2000) deﬁned spoilage as “The change in
characteristics in a food, making it no longer acceptable”, spoilage in foods can
be considered subjective. Different factors, such as insects, dehydration and
environment conditions, cause spoilage; but by far, one of the primary causes
comes from microbial activity (Adams and Moss, 2000). Fungi have been
identiﬁed as the major agents of postharvest fresh chestnut decay and rot
(Jerimini et al., 2006; Jian et al., 2002; Montealegre and Gonzalez, 1986; Miller
G., 2003; Paglietta and Bonous, 1979; Rutter et al., 1990; Vettraino et al., 2005;
Vossen, 2000; Washington et al., 1997). A broad number of organisms have
been identiﬁed that cause chestnut decay, colonizing the product after harvest.

Few of all these organisms, have been characterized as true plant
pathogens, capable of initiating infection prior to harvest. One true plant pathogen
Sclerotinia pseudotuberosa Rehm (syn. Cibon'a batschiana Zopf., anamorph
Rhacodiella castaneae Bainier), is responsible for chestnut black rot and one of
the most important postharvest diseases of acorns (Quercus spp.) and chestnuts,
especially in Europe (Vettraino et al., 2005; Washington et al., 1997). Infection
usually occurs when chestnuts fall to the ground during harvest and rapidly

become contaminated with ascospores secreted by fungal apothecia, previously

13

formed by dormant sclerotia lying on the ground (Agrios, 2005; Vettraino et al.,
2005). However, studies by Delatour and Morelet (1979) and Vettraino et al.,
(2005) have also found infected chestnuts attached to the tree, and have
identiﬁed the pathogen present in different plant parts, suggesting that the
pathogen may occur symptomatically and asymptomatically as an endophyte.
Another uncertain but possible real pathogen, which affects chestnuts is
Phomopsis castanea Sacc., causing phomopsis chestnut rot. This organism has
been predominantly reported in Australia and Chile and was recently identiﬁed as
an endophyte in chestnuts. Although the disease cycle is still unclear, this fungus
colonizes nuts via the peduncle and hilum (Appendix C). The disease can also be
found in both chestnuts attached to the tree, and those in storage, but it appears
that natural infection occurs in the ﬁeld (Washington et al., 1998). Partial control
of the disease can be achieved by postharvest management practices including
low temperature storage and controlled atmosphere storage following an initial
period in a controlled carbon monoxide atmosphere (Washington et al., 1998).
Natural infection occurs in the ﬁeld before harvest, since this disease can be
found in chestnuts both before and immediately after harvest. Therefore, control
measures are needed to restrict disease development in the ﬁeld. Few report the
use of fungicides for control of Phomopsis chestnut rot or related diseases in
chestnuts. Montealegre (1984) indicated that captan, dodine or a combination of
benomyl and mancozeb showed good control of P. castanea in vitro. Washington
et al. (1998), reported that benomyl, imazalil, prochloraz and propiconazole were

most effective in vitro against mycelial growth in Australia.

14

Opportunistic fungi involved in chestnut mold and rot after harvest include
Phomopsis endogena Speg. (cif. Phoma endogena Speg.), Phomopsis viterbenis
Camici, Diplodia castanea Prill and Del., Alteman'a sp, Aspergillus sp., Penicillium
spp., Botrytis cinerea Pers., Gloespon'um sp., Fusan'um spp., Dothiorella sp.,
Cytodiplospora castanea Oud., Pestalotia spp., Diplodia spp., Acrospeira
mirabilis Berk. and Broome and Rhizopus spp (Breisch, 1993; McCarter et al.,
1980; Montealegre and Gonzalez, 1986; Paglietta and Bonous, 1979; Pratella,
1994; Puttoo et al., 1988; Ride and Gudin, 1960; Washington et al., 1997; Wright,
1 960).

Peeled chestnuts, considered to be a fresh-processed (cut) product (IFPA,
2006) can be affected by the same microorganisms as the fresh product, but
other opportunistic organisms may also play an important role in their spoilage.
By removing the shell and pellicle, which are considered to be a weak but still
natural physical barriers that protect the kernel, contamination with pathogens,
opportunistic organisms and water loss can be facilitated, affecting their ﬁnal
quality and safety (Cantwell, 1995; Mencarelli, 2001). Therefore, consideration
should be given to removing the shell and other plant parts, since natural
products derived from plant parts may contribute to pathogen resistance and
colonization by opportunistic organisms (Field et al., 2006). Such compounds,
involved in protection from decay have not yet been isolated from chestnut
kernels, however evidence suggests that undetermined substances in the shell
and pellicle, may protect the kernel. These presumptive compounds may protect
the chestnuts from decay, especially when the chestnuts are on the tree or stored

fresh (Appendix E).

15

Even though Michigan peeled chestnuts have experienced microbial
spoilage during cold storage (4 °C), there have never been reports that indicate
similar spoilage problems in peeled chestnuts elsewhere. However, this problem
is not unique because similar spoilage occurs with other vegetables and fruits,
mainly after peeling, packaging or processing (Brightwell et al., 2007; Guerzoni et
al., 1996; James et al., 2005; N9, 2007; Ragaert et al., 2006; Tournas, 2005;
Zagory, 1998). Zagory (1998) and Tournas (2005) indicated that after harvest, a
wide range of economically important vegetables and speciﬁcally fresh-cut
products, are often spoiled by a wide variety of microorganisms including many
bacterial (Curtobacten'um, Rahnella, Enivinia, Pseudomonas spp., among others)
and several fungal species. Of those commodities evaluated, spoilage increases
signiﬁcantly when the product is injured or the skin, which acts as a physical
barrier has been damaged or removed. Packaged sliced onions, shredded mixed
lettuce and other commodities under ambient conditions and modiﬁed
atmosphere, can become colonized by diverse spoilage microorganisms,
including various yeasts as Pichia fennentans, Cryptococcus Iaurentii, and
Candida spp., among others (Liu and Li, 2006; Ragaert et al., 2006).

Microbial populations typically increase from harvest, through processing,
and cool storage, with the extent of growth impacting the shelf-life of the product.
This can lead to a series of problems in both fresh and processed products,
which includes several types of external molds, rots, internal decay and severe
spoilage (Hammer, 1949; Tournas, 2005; Washington et al. 1997; McCarter et

al., 1980; Zagory, 1998). These microbial changes, bring about undesirable

16

qualities and economical loss, thus conﬁrming the importance of controlling
microbial growth during production, harvest, processing, and during storage
(Tournas, 2005; Zagory, 1998).

Food safety concerns

A critical factor in developing any industry is the ability to offer a safe high
quality product. Chestnuts are not only subject to microbial decay, but without
proper care, safety of the product may be subsequently compromised by
accidental contamination with food-borne pathogens and an increase in endemic
organisms or compounds. These organisms and compounds may cause
immediate or long-term problems and diseases as well as intoxication of the
consumers. Food has historically been associated with the transmission of
diseases (CDC, 2008).

The World Health Organization (WHO, 1992) states: “Even today, despite
the increase of knowledge, foodbome disease is perhaps the most widespread
health problem in the contemporary wor1d and an important cause of reduced
economic productivity’. The Centers for Disease Control and Prevention (CDC)
estimate that annually in the United States, at least 76 million cases of foodbome
illness occur, more than 300,000 people are hospitalized and 5,000 die (CDC,
2008). In the United States, during 1998 to 2002, a total of 6,647 outbreaks of
foodborne disease were reported, causing approximately 128,370 people to
become ill (CDC, 2008). In 33 percent of the cases, the etiology was identiﬁed
with bacterial pathogens causing the largest percentage of outbreaks (55
percent). Among bacterial pathogens, Salmonella enterica serotype Enteritidis

and Listeria monocytogenes accounted for the largest number of outbreaks and

17

the majority of deaths. In addition, multistate outbreaks caused by contaminated
fresh produce, such as spinach and lettuce, mainly caused by Escherichia coli
O157:H7 have remained prominent. Viral pathogens, predominantly norovirus,
caused 33 percent of outbreaks and increasing. Chemical agents, like
mycotoxins, and parasites caused 10 and 1 percent of the outbreaks,
respectively (CDC, 2006; CDC, 2008).

Thus far no foodbome disease outbreaks have been traced to chestnuts in
the United States. Nevertheless, this cannot be discarded due to the fact that
since 1998, more than 18 foodbome disease outbreaks in fruits and other nuts
have been reported each year. Of those, in 2006, Salmonella serotype
Tennessee and Salmonella serotype Thompson, together caused 107 cases of
salmonellosis in Vermont and South Carolina, mainly from consumption of mixed
nuts and peanuts (CDC, 2006). Furthermore, in 2004 an outbreak of
salmonellosis has been linked to consumption of contaminated raw almonds, and
hundreds of consumers across the United States may have been sickened. The
recall was expanded to include millions of pounds of raw California almonds sold
worldwide, signiﬁcantly affecting the reputations and economy of this almond
industry (CDC, 2006).

The number of outbreaks due to the presence mycotoxins is low, this
natural toxins impact the safety of the ﬁnal product, especially when long-term
health effects are considered. Concerns over mycotoxins contamination has
continued to climb in recent years. Recent recalls in several industries, including
the grain and pet food industry in the United States have raised questions over

ingredients sourcing and cross-contamination of the food chain. This afﬁrms that

18

the economic impact of mycotoxins in feeds is in the hundreds of millions of
dollars, including reduced production, monitoring, managing and control (CDC,
2008). Considerations must be taken in account, because diverse fungi colonize
chestnuts and mycotoxins have been identiﬁed, in the fresh product during
storage and retail sales (Overy et al. 2003).

New technologies such as modiﬁed atmosphere packaging (MAP) and
other innovative safe postharvest treatments can reduce decay and spoilage
organisms. Even though the food may look and smell appropriately, the product
may not be safe to eat (Rajkowski and Baldwin, 2003). In diverse fresh-cut
products such melons and other fruits that have been packaged under MAP, an
increase in CO; and decrease in 02 has been observed, which favor Clostridium
botulinum growth and toxin formation, making these products unsafe for
consumption (Enfors and Molin, 1978). When minimally processed and packaged
foods are stored under refrigerated conditions (4 - 7° C) Salmonella, Aeromonas,
Yersinia, Vibrio cholerae, E. coli O157:H7, and other foodbome pathogens can
grow to undesirable levels (Novak et al., 2001). All of these food safety issues
must be considered as possible safety concerns for fresh chestnuts, and could be
especially devastating in the case of processed products, like vacuum-packaged,
peeled, frozen chestnuts.

Due to these concerns, when any food product is being distributed,
including fresh and peeled chestnuts, preventive measurements must be taken to

assure that the product is safe to consume (CDC, 2006).

19

Sanitizers, postharvest and post processing treatments

VIﬁth increased global trade and seasonality of agricultural products, fresh
agricultural commodities are being transported over vast distances and stored for
prolonged periods. Thus, effective cold-chain and product management practices
are required to ensure that fresh products maintain their premium quality and
safety. The fresh produce industry must develop several of the impact of
postharvest treatments, sanitizers and practices that contribute to produce quality
and safety (Kader, 2002).

Sapers (2003) stated that, “Generally, it is more difﬁcult to decontaminate
products than it is to avoid contamination. Therefore, pre- and postharvest
interventions that reduce risk of contamination will make the job of sanitizing the
produce easier”. Due to this situation, the US. Food and Drug Administration
(FDA) published a “Guide to Minimize Microbial Food Safety Hazards for Fresh
Fruits and Vegetables” in October 1998, which describes the important steps
needed to remove ﬁeld contaminant (dirt, twigs and more), chemical residues,
and microorganisms responsible for quality loss, safety concerns and decay. This
was a general, commonsense guide because, at that time, there was very little
speciﬁc research to provide more concrete advice. In response to the continuing
problems in particular sectors of the fresh produce industry, FDA has issued
several warning letters and initiatives. In January 2004, FDA and CDC met with
produce industry leaders to discuss numerous foodborne illness outbreaks
associated with produce. Industry representatives agreed to take the lead on

developing commodity-speciﬁc Good Agricultural Practices (GAPs) that would

20

provide additional guidelines tailored to individual commodities that had been
implicated in recent foodborne illness outbreaks. In April 2006, in reaction to E.
coli O157:H7 outbreaks on lettuce and spinach, the produce industry published
its “Commodity Speciﬁc Food Safety Guidelines for the Lettuce and Leafy Greens
Supply Chain.” Nevertheless, until recently, little information was available
concerning the efﬁcacy of washing and sanitizing treatments (Sapers, 2003).

Historically, more is known about chemicals, such as fungicides, but
consumer attitudes, safety concerns, regulations, and health and environmental
issues regarding their use, inhibit their implementation. Nevertheless, these
broadly used are efﬁcient in reducing decay organisms (Gamage et al., 1997;
Kader, 2002; Kicast, 1995; Narayanasamy, 2006; Sapers, 2003; Saroj et al.,
2006; Smith and Pillai, 2004). Many less toxic have since been investigated
including hydrogen peroxide, diverse acids, ozone, chlorine dioxide (gas and
solution), hypochlorite, organic acids, natamycin, heat and more (Beuchat 1998;
Block 1991; Brackett 1999; Crowe et al., 2005; Palou et al., 2002; Panagou et al.,
2005; Perrera and Karunaratne 2001; Pierre et al., 2006, Sapers 2003; Suslow
2002; Wade et al., 2003). These compounds can be used to enhance safety,
and shelf life of most type of fresh produce and including blueberries, citrus,
sprouts, meat, leafy greens and many others. Several salts, oils and biocontrol
agents that efﬁciently reduce decay also have been evaluated (Feng and Zheng,
2006, Kader, 2002; Mari et al., 2003; Mills et al., 2004; Porat et al., 2002; Sapers,
2003; VIﬁllis et al., 2007).

Incoming produce is typically delivered to a storage or distribution facility.

After delivery, it is immersed in clean water containing an appropriate

21

concentration of a speciﬁc chemical sanitizer, or treated by other means (e.g.
irradiation) with the ﬁnal objective being the elimination of pathogens and the
reduction of other organisms to an acceptable level (Kader, 2002; Sapers, 2003).
The advantages, limitations, usage restrictions of a number of agents used to

sanitize horticultural commodities and diverse foods are described in Table 1.

22

 

 

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Sapers (2003) indicated that washing and sanitizing produce and
equipment generally does not reduce the microbial populations greater than 2-
logs (99 percent). Although these reductions may improve products shelf-life and
quality, the possibility of human pathogen survival can not be excluded.
Furthermore, diverse factors such as microbial adherence, formation of bioﬂlms,
and penetration of the microorganisms may compromise the efﬁcacy of certain
compounds. New washing technologies are being constantly evaluated and are
needed to overcome these deﬁciencies. The new treatments must be superior in
efﬁcacy, but also must be safe and affordable to apply. Innovating technologies,
such as food irradiation and heat treatments, which may be capable of reducing
pathogens and other microorganisms (pasteurization treatments), greater than 5-
logs might bring large improvements in the microbiological quality and safety of
produce (Niemira, 2003).

Various chemical treatments have been evaluated for chestnuts in
attempts to reduce mold and decay during fresh chestnut storage. Among the
evaluated treatments, 95 percent alcohol, 1 percent copper sulfate, boric acid,
0.4 percent calcium chloride, formaldehyde, benzoic acid, sulfur dust and others
were unsuccessful. Furthermore, these experiments indicated that certain
treatments such as boric acid and formaldehyde, might also compromise the
quality of the chestnuts by hardening the kernel (Hammer, 1949; Tan et al.,
2007). Other materials exhibiting potential for decay control include 30 ppm
iodine, 250 ppm carbendazim, 100-200 ppm sodium hypochlorite, 100 ppm

peracetic acid (Morris, 2006; Panagou et al., 2005), 50 ppm natamycin, 1 percent

28

sorbic acid, 1 percent propionic acid, hot water dip at 90 °C for 10 minutes
(Panagou et al, 2005) and immersion in certain fungicides like 2,6-dichloro-4-
nitroaniline (Botran) (Wells, 1980). Water immersion at 15 °C for 15 minutes, 45-
48 °C for 15 minutes and 52 °C for 5, 15, 30 or 60 min signiﬁcantly reduced the
insect damage and the presence of larvae of Cydia splendana, Curculio elephas
and Curculio sayi during storage (Jerimini, et al., 2006; Sieber et al., 2007; Wells,
1980).

Even though various treatments may reduce microbial populations and
insects in chestnuts water immersion at 52 °C warm bath for 60 minutes, may
affect the ﬁnal quality of the product, by decreasing soluble sugars and increasing
starch during storage (Wells, 1980). Other options may be available to treat
chestnuts after harvest, however there is still a need to ﬁnd feasible safe
alternative approaches to control decay of fresh chestnuts, during storage
(Sapers, 2003).

Limited work has been done on strategies to control postprocessing
spoilage of peeled chestnuts. However, many treatments used broadly for other
food commodities may be useful in enhancing the microbial shelf-life of chestnuts

(Sapers, 2003).

Objectives of this study

Several studies have identiﬁed the organisms that colonize chestnuts
(Jerimini et al., 2006; Jian et al., 2002; Montealegre and Gonzalez, 1986; Miller
G., 2003; Paglietta and Bonous, 1979; Rutter et al., 1990; Vettraino et al., 2005;

Vossen, 2000; Washington et al., 1997). However, concerns regarding local

29

postharvest molds and decay in fresh chestnuts, as well as postprocessing
microbial spoilage in peeled chestnuts still remain. Studying the organisms that
colonize chestnuts and their interaction with each of the products should help us
understand the behavior of these organisms before harvest, during storage and
after processing. These studies will provide powerful tools to increase shelf life,
leading to improved marketing of attractive, safe high quality chestnuts.
Therefore, to help determine the impact of various microorganisms in the shelf
life of fresh and peeled chestnuts, two objectives were set:

1. Determine the population of microorganisms and their impact on shell
mold and kernel decay of fresh chestnuts in Michigan.

2. Determine the population of microorganisms and their impact on
mechanically peeled chestnuts in Michigan.

Routine postharvest application of sanitizers, such as chlorine to
chestnuts, does not appear to provide efﬁcient control of the previously
mentioned postharvest or postprocessing problems, leading to signiﬁcant
economic and quality losses. In reaction to this concern, two objectives were
designed to identify more efﬁcient microbial reduction strategies to minimize
postharvest decay of chestnuts:

1. Evaluate the efﬁcacy of postharvest treatments to reduce microbial
decay of fresh chestnuts.

2. Evaluate the efﬁcacy of postprocessing treatments to reduce microbial
spoilage of peeled chestnuts.

With answers to these questions, I hope to help the young chestnut

industry in Michigan provide high-quality chestnuts in future years.

30

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42

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43

CHAPTER 2
SHELL MOLD AND KERNEL DECAY OF FRESH CHESTNUTS IN MICHIGAN
Abstract
Chestnut is a relatively new crop for Michigan and post-harvest loss due to
decay has been problematic as production has increased each year. In 2007,
more than 25 percent of the nuts were lost to postharvest decay, equivalent to
approximately 5,300 kg of fresh product. To determine the organisms responsible
for decay, a microbiological survey was performed in 2006 and 2007 to identify
microorganisms involved in postharvest shell mold and internal kernel decay of
chestnuts. Filamentous fungi including P. expansum, P. griseofulvum, P.
chysogenum, Coniophora puteana, Acrosperia mirabilis Acrospaeria mirabilis,
Botryosphaeria ribis, Sclerotinia sclerotiorum, Botryotinia fuckeliana (Anamorph
Botrytis cinerea) and Gibberella sp. (Anamorph Fusarium sp.) were the
predominant microorganisms that negatively impacted fresh chestnuts.
Populations of microorganisms, including these fungi, varied between the farms,
harvesting method and chestnut part. Overall, chestnuts harvested from the
orchard ﬂoor were signiﬁcantly (p < 0.05) more contaminated than chestnuts
harvested directly from the tree, by more than 2-Iog CFU/g. In addition, a
signiﬁcant difference (p < 0.05) in the microbial population was seen between
chestnuts submitted by different growers, with average count ranges of fungi,
mesophilic aerobic bacteria (MAB) an yeast equal to 4.75-, 4.59- and 4.75-log

CF U/g subsequently.

44

Introduction

Chestnut (Castanea spp.) is considered to be the most popular nut-bearing
tree among the Mediterranean countries of Europe and many countries in Asia
with new production now starting in Australia, New Zealand and Chile (Fulbright
and Mandujano, 2000; Grau and France, 1999; Klinac, Seelye, and Nguyen,
1999; Ridley, 1999; Mencarelli, 2001). The increased market for chestnuts is
partially due to their nutritional value, low lipid content and antioxidant properties,
(Anagnostakis and Devin, 1999; Biomhoff, Carisen, Andersen, and Jacobs, 2006;
Gao, Lin, and Xiao, 2008). In the United States chestnuts are rare. People are
more likely to be familiar with the unrelated poisonous horse chestnut (Aesculus
spp.) (Fulbright and Mandujano, 2000) than with any of the edible, sweet
chestnut species (C. dentata Borkh., C. mollissima BI., C crenata Sieb and Zucc.,
C. sativa Mill., C. seguinii Dode, C. pumila Mill, C. henryi Rehd. and ers.)
(Anagnostakis, Gordon, and Hebard, 1998; Fulbright and Mandujano, 2000;
Miller, 2003). This is partially due to chestnut blight (Cryphonectria parasitica
Murril and Barr), which virtually eliminated the once-widespread American
chestnut during the ﬁrst half of the twentieth century (C. dentate) (Anagnostakis,
1992). Nevertheless, commercial orchards have been established in different
locations in the United States, and have been increasing over the years (Fulbright
and Mandujano, 2000; Vossen, 2000). The United States, has at least of 2,500
acres of chestnuts tree plantations. Of these, approximately 1,500 acres of young
(less than 10 years) chestnut trees are distributed among Michigan, California,

Oregon, Washington and Ohio (Fulbright, 2008; Vossen, 2000). Commercial

45

plantations are primarily composed of the cultivar ‘Colossal’ (European-Japanese
hybrid = C. sativa x C. crenata). Other cultivars planted are composed
predominately of Chinese chestnuts due to the naturally occurring chestnut blight
resistance in this species (Miller, 2003; Fulbright and Mandujano, 2000; Vossen,
2000). Mature seedling Chinese chestnut trees (non-grafted trees) also have
been planted in Midwest and eastern states with some orchards of seedling
European chestnuts.

In the past ten years, domestic chestnut production has increased
signiﬁcantly. North American chestnuts are typically harvested from late-
September to the ﬁrst week of November. Vlﬁth most chestnuts sold from
Thanksgiving through Christmas to specialty ethnic markets, retail stores, food
processors, restaurants, holiday festivals, farmers’ markets and individual
consumers. In Michigan, fresh chestnuts and frozen, peeled chestnuts comprise
up to 60 and 30 percent of outlet sales, respectively. The remainder is sold
dehydrated as ﬂour, breading and a new product called chestnut slices
(Blackwell, 2006; Fulbright D. W., 2008).

Since freshness and microbial quality are factors in chestnut marketing,
domestic production has a deﬁnite advantage over imported chestnuts. In
Michigan, production is limited primarily due to the young nature of the industry
and the microbial decay that occurs during storage. Postharvest decay of
chestnuts causes severe losses, leading to completely unmarketable chestnuts.
Several molds are known to cause severe postharvest decay and disease
problems in chestnuts worldwide. Among these, Penicillium sp., Aspergillus sp.,

Fusarium sp., Phomopsis castanea, Acrospeira mirabilis, and Sclerotinia

46

pseudotuberosa (syn. Ciboria batschiana, S. batschiana; anamorphic from
Rhacodiella castanea, syn. Myrioconium castanea) have been repeatedly
identiﬁed in France, Italy, Australia, Chile and the United States (Ellis and Ellis,
1985; Jerimini, et al., 2006; Montealegre, and Gonzalez, 1986; Paglietta and
Bonous, 1979; Ride and Gudin 1960; Sieber et al., 2007; Vettraino et al., 2005;
Washington et al. 1997; Wright, 1960.

Three studies were design to better understand the impact and possible
effect of microorganisms involved in postharvest chestnut shell mold and internal
kernel decay in Michigan. The ﬁrst study was set up on one farm, with the
purpose of evaluating the effect of different harvesting methods on microbial
population. The second study qualitatively assessed the microbial quality (shell
mold severity and incidence of kernel decay) from seven different farms. The
third study identiﬁed microorganisms involved in shell mold and kernel decay on

chestnuts collected from at least seven different farms.

Material and Methods

Collection and storage of chestnut samples

For the ﬁrst study, were the objective was to evaluate the effect of different
harvesting methods on microbial population, chestnuts of the cultivar 'Colossal’
were collected from one farm in Livingston County, MI during the 2006 and 2007
growing season. Mature chestnuts were sampled using a stratiﬁed sampling
method with two levels. The ﬁrst level consisted of nine chestnut samples (~0.9

kg each) taken from the ground after less than 24 hours of contact. The second

47

level was composed of nine mature chestnut samples (~0.9 kg each) hand-
harvested from different tree heights, in mature burs. Each sample was placed in
a plastic bag and transported in a portable cooler to the storage facility. Before
long-term storage, each chestnut sample was transferred to a mesh bag and
randomly stored in a cooler at 4°C, located in Michigan State University (East
Lansing, Michigan). Samples were separately placed on racks to prevent cross
contamination. Microbial populations on chestnuts shells and kernels from two
consecutive growing seasons (2006 and 2007) after harvest were assessed, and
at 30, 60, 90 and 120 days of storage.

The second study consisted in evaluating differences between shell mold
severity and kernel decay incidence (microbial quality), among farms in Michigan,
three 0.77 kg samples of 'Colossal' chestnuts per farm were randomly collected
before'storage during the 2007-growing season. Each sample was randomly
stored in mesh bags, as described above.

In addition, to engage the third study, a total of 200 individual ‘Colossal’
chestnuts from the 2006 and 2007 growing seasons, collected from at least
seven different farms were randomly sampled twice during storage to identify the
organisms that colonized the chestnuts. From this group, chestnuts showing shell
mold or kernel decay symptoms were sorted for isolation and identiﬁcation of the
causal agents. Before identiﬁcation, the observed symptoms were described,

photographed and classiﬁed.

48

Microbial populations — First and third study

Shell

To determine the microorganisms found on shells, 25 g chestnut samples
were placed in a sterile stomacher bag (Whirl-Pak, NASCO, International Inc.),
diluted 1:5 in phosphate buffer solution (PBS) (pH = 7.4) and agitated for 1 min
(2900/3500 rpm) in a pulsiﬁer (Filtaﬂex Ltd., Almonte, Ont., Canada). After serial
dilution, 100 pl aliquots were inoculated onto trypticase soy agar (Becton and
Dikinson, Md, USA) containing 0.6% BactoTM yeast extract (Becton and Dikinson)
and 100 ppm of cycloheximide (Sigma-Aldrich, Mo, USA) for quantiﬁcation of
mesophilic aerobic bacteria (MAB) and onto potato dextrose agar (Becton and
Dikinson) containing 20 ppm streptomycin (Sigma-Aldrich) and 50 ppm ampicillin
(Sigma-Aldrich) for enumeration of yeasts and molds. The populations of MAB
were determined after 48 h at 25° C (i 3° C). Mold and yeast plates, were

counted after 72 h of incubation at 25° C (:I: 3° C) (APHA, 2001).

Kernel

ldentically stored chestnuts from the same samples were ﬂame sterilized
after dipping in 99% ethanol. After manually removing the shell, each sample was
weighed and placed in a sterile stomacher bag (VVhirI-Pak), diluted 1:5 in PBS
and homogenized for 1 min (normal speed) in a stomacher Model 400 (Seward
Lab. System, England). The suspension was serially diluted, and quantitatively

examined for MAB, yeast and molds as previously described (APHA, 2001).

49

Microbial mold severity and kernel decay incidence analysis - Second

study

A sub-sample consisting of 3 randomly chosen stored chestnuts collected
from each of seven farms was assessed for severity of shell mold and incidence
of kernel decay after harvest and after 30 and 60 days of storage. Shell mold
severity was calculated as the mean number of a 4-point visual qualitative
measurement of decay, where 0 = 0 % decay, 1 = 1-25 % decay, 2 = 26-50 %
decay, 3 = 51-75 % decay and 4 = 76-100 % decay. Incidence of decayed
kernels was calculated as the mean number of chestnuts showing symptoms of

decay.

Identiﬁcation of microorganisms - Third study

All of the microorganisms isolated from symptomatically chestnuts,
containing shell mold or kernel decay were picked and subsequently puriﬁed after
isolation. MAB and molds were identiﬁed sequencing rDNA subunits (White,
Bruns, Lee, and Taylor, 1990).

Genomic DNA was extracted from molds using the DNA QiAquick DNA
extraction kit (Qiagen, Maryland, USA) as recommended by the manufacturer. A
0.5 to 1 mg sample of mycelia taken from ﬁve- to seven-day-old molds grown on
potato dextrose agar, was immediately ground in a sterile mortar and pestle
containing 500 pl of cetyltrimethylammonium bromide (CTAB) buffer pH 8.3
(Qiagen, Maryland, USA). After these the DNA was extracted followed the
procedure proposed by Hamelin et al. (2000). The region including the two

spacers (ITS4 and ITS6) was ampliﬁed. A total volume of 25 pl for each reaction

50

contained 60 ng of DNA, 23 pl of AFLPT'“ ampliﬁcation core mix (Applied
Biosystems, Foster City, Ca, USA), 2.5 units of AmpliTaq Gold DNA polymerase
(Applied Biosystems) and 10 pM of each primer. The primers used, ITS4 (5'-
TCCTCCGCTTATTGATATGC-S') (White et al., 1990) and ITS6 (5'-
GAAGGTGAAGTCGTAACAAGG-3’) (Cooke et al., 2000) are complementary to
the ﬁnal portion of 18S rDNA adjacent to ITS3 and to the initial portion of 25S
rDNA adjacent to ITSZ, respectively (Cooke et al., 2000; White et al., 1990).

Bacterial genomic DNA was extracted using the protocol proposed by
Jacobs et al. (2008). The region including the two spacers (UF LP and URPL) was
ampliﬁed using the method of LiPuma et al. (1999). A total volume of 25 pl for
each reaction contained 60 ng of DNA, 23 pl of AFLPTM ampliﬁcation core mix
(Applied Biosystems, Foster City, Ca, USA), 2.5 units of AmpliTaq Gold DNA
polymerase (Applied Biosystems) and 10 pM of each primer. The primers UFPL
(5'-AGTTTGATCCTGGCTCAG-3') and URPL (5'-GGTTACC'ITGTTACGACTT-
3') were used for targeting the 16S rDNA region from the kingdom Procaryotae
(bacteria) (LiPuma et al., 1999).

For molds and bacteria the extracted DNA region was ampliﬁed in a 2720
thermal cycler (Applied Biosystems). The program included a cycle at 95 °C for 2
min, 30 cycles of 95 °C for 30 s, 55 °C for 30 s and 71 °C for 1 min, and a ﬁnal
elongation at 71 °C for 10 min. The PCR product was placed in wells on an
agarose gel (1.25 %), and electrophoresed for 3 hours at 70 volts. After
electrophoresis, the ampliﬁed region was puriﬁed using a QlAquick PCR

Puriﬁcation kit (Qiagen, Maryland, USA). The PCR product was sequenced in

51

both directions using moderate throughput sequencing (Research Technology
and Support Facility, MSU, East Lansing, MI, USA). Sequences were deposited
in the Lasergene software (DNA Star Inc., Madison, WI., USA) and used as
queries for similarity searches in the NCBI nucleotide database (NCBI, 2008).
Species reported had a 98 to 100 % match in both directions.

Yeasts were inoculated on potato dextrose agar, incubated for 72 h at 25°
C (:t 3° C) and then sent to the DCPHA (MSU, East Lansing, Mi., USA) for further
identiﬁcation. Yeasts were identiﬁed using the API 200 yeast identiﬁcation
system together with microscopic morphology determinations (Michigan State
University Diagnostic Center for Population and Animal Health (DCPHA)). This
computer-assisted system for rapid identiﬁcation of yeast provides results
comparable to those obtained of conventional morphological methodologies

(Land et al., 1979).

Statistical analysis

One-factor and repeated measurement design with analysis of variance
(ANOVA) were done on all microbial count and microbial qualitative assessment
data obtained from fresh chestnuts. Since all chestnuts were randomly assigned
to the different treatment conditions, sphericity can be assumed making
multivariate as well as degrees of freedom corrections unnecessary (Keselman,
Algina, and Kowalchuk, 2001; Ott and Longnecker, 2001). Multiple analysis of
variance (MANOVA) was also used to determine differences between the
microbial populations in chestnuts from different farms. Signiﬁcance between

means was determined using the Tukey post-hoc multiple comparisons of means

52

test at the 95% family-wise conﬁdence level (p=0.05). Calculations were
performed using the statistical package “R: A language and environment for
statistical computing” (R Development Core Team, 2007) (Ott and Longnecker,

2001)

Results

Shell mold symptoms

The harvested chestnuts were colonized by a broad range of
microorganisms (Table 2), not all of which had the potential to cause shell mold.
Fungi that infested and caused shell mold on fresh chestnuts were mainly
composed of three species of Penicillium (P. expansum, P. griseofulvum and P.
chysogenum) and Coniophora puteana. Dark-green to black spots with white
mycelia were observed on chestnuts colonized by Penicillium species. These
spots begin on the hilum (Figure 1—a and Appendix C). Once the mold grew, a
mantle of white mycelia containing dark-green, blue, bluish-green or olive-green
completely covered the shell (Figure 1-b). In extreme conditions, the fungi
softened and discolored the shell tissue, subsequently affecting the quality of the
fresh product (Figure 1-c). Symptoms from Penicillium were ﬁrst observed after
harvest, and rapidly developed during storage. Penicillium species belong to the
phylum Deutoromycota, producing green, bluish-green or olive-green spores
asexually. Various species cause blue mold rots and green mold rots. They are
the most common and usually the most destructive of all postharvest diseases,
affecting various fruits and vegetables, including commodities of importance like

apples and citrus (Agrios, 2005). In addition to the losses caused, the fungus also

53

produces several mycotoxins, such as patulin and zearalenone (Adams & Moss,
2000).

Coniophora puteana, rarely infected the shell, but when present, the
organism developed in pockets of white web-like mycelia around several
chestnuts (Figure 1-d), which contained apparent high moisture content. After
manually removing the mold, the appearance and quality of the chestnuts were
not affected. This fungus belongs to the phylum Basidiomycota. It has never been
reported affecting chestnuts or other commodities, but it can be found affecting
indoor wood structures and stored wood, causing wet brown rot, signiﬁcant decay

and economical damage (Schmidt, 2007).

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58

 

 

 

 

 

 

Figure 1. Chestnuts showing shell mold due to a-c) Penicillium spp. (P. griseofulvum, P.
expansum, P. chysogenum) and d) Coniophora puteana. Image in this thesis is presented
in color.

Kernel decay symptoms

Microorganisms infecting chestnut kernels primarily included Penicillium
spp. (P. griseofulvum, P. expansum, P. chysogenum), Acrospaen'a mirabilis,
Botryosphaen'a ribis, Sclerotinia sclerotiorum, Botryotinia fuckeliana (Anamorph
Botrytis cinerea) and Gibberella sp. (Anamorph Fusan'um sp.). Penicillium spp.
(P. griseofulvum, P. expansum, P. chysogenum) produced white mycelia and
dark-green, blue, bluish-green or olive-green spores (Figure 2-a). That
sometimes penetrated deep into the kernel, resulting in extensive kernel rotting
and complete kernel decay (Figure 2-b). Acrospaen'a mirabilis, appeared as dark-

brown spots (conidia), ﬁlling the space between the kernel cotyledons and kernel

59

cracks. Whitish, web-like mycelia developed around these spots. Brown necrotic
spots were observed around the colonies, which sometimes enlarged, turned
light brown, and ﬁnally dark-brown. The infection sometimes penetrated deep into
the kernel, softening the tissue, and resulting in opaque brownish kernel decay.
(Figure 2-c). This fungus causes considerable losses in chestnuts during storage,
belonging to the phylum Deuteromycota. It lacks fruiting bodies, producing
conidia asexually. Conidia are produced coiled, with 2 pale-cells and 1 dark-
brown, verrucose, globose terminal cell (20-30 u m diameter) (Ellis & Ellis,
1985)

Botryosphaen'a ribis and Sclerotinia sclerotiorum were isolated from only a
few kernels that presented an extreme dark-black decay (Figure 2-d). In some
situations, infected tissue became hard and completely mummiﬁed.
Botryosphaen‘a n'bis and Sclerotinia sclerotiorum, belong to the Phylum
Ascomycota. Botryosphaeﬁa ribis produces conidia in pycnidia, while the fungus
Sclerotinia sclerotiorum overwinters as sclerotia on or within infected tissues and
in the ground. In spring or early summer sclerotia germinate and produce
apothecia, which contain asci, producing ascospores (Agrios, 2005).

Botryotinia fuckeliana (Anamorph Botrytis cinerea) was isolated from
grayish, completely decayed kernels. The mycelium appeared to rapidly invade
the whole kernel, which became covered with a whitish-gray, cobweb-like mold.
Infected kernels became soft, watery and acquired a dark gray to black coloration
(Figure 2-e). Depending in its sexual stage, this fungus belongs to the phylum
Ascomycota or Deuteromycota. It causes gray molds or rots of fruits and

vegetables, both in the ﬁeld and during storage. It produces abundandt gray

60

mycelium and long branches of conidiophores, which contain rounded apical cells
bearing folorless or gray, one-celled, ovoid conidia (Agrios, 2005)

Gibberella sp. (Anamorph Fusarium sp.) caused white mold decay.
Affected kernels appeared dry, whitish to light brown and extremely pale. A
whitish web-like mycelium could be observed around decayed tissue, usually in
the cracks, (Figure 2-f). Depending in its sexual stage, this fungus belongs to the
phylum Ascomycota or Deuteromycota. The fungus usually produces colorless
mycelia, at ﬁrst, but with age it becomes cream-colored, pale yellow, pale pink, or
purplish. It produces three kinds of asexual spores, microconidia, macroconidia
and chlamydospores, but also produced sexual spores, throught perithecia,
during its sexual phase (Agrios, 2005). In this study, this microorganism was
usually mistaken for early infection by Penicillium spp., and usually both were
present. In addition, the fungus also produces several mycotoxins, such as DON

and zearalenone (Adams & Moss, 2000).

61

 

 

 

 

 

 

 

 

:igure 2. Chestnuts showing kernel decay due to a-b) Penicillium spp. (P. griseofulvum, P.
expansum, P. chysogenum), c) Acrospaeria mirabilis d) Botryosphaeria ribis and
Sclerotinia sclerotiorum, e) Botryotinia fuckeliana (Anamorph Botrytis cinerea), f)
Glbberella sp. (Anamorph Fusarium sp.). Image in this thesis is presented in color.

Inﬂuence of chestnut harvesting method on microbial populations
Non-decaying chestnuts yielded low populations of MAB, molds an yeasts
on the shells and in kernels when collected on day 0, from trees and the ground.
The microbial population signiﬁcantly increased during storage (F (3,4) = 176.61,
MSE = 5.81, p < 0.01) (Figures 3, 4 and 5). These populations ranged from 1.8 to
4.61 CFU/g log, depending on the portion of the chestnut examined (shell or
kernel) and the harvesting method (ground or tree). With the shell yielding
signiﬁcantly higher (F (1,4) = 324.09, MSE = 1.3 p < 0.01) microbial populations

compared to the kernel.

62

In general, populations of molds and MAB tended to increase during 60 to
90 days of storage, regardless of the harvest procedure (Figures 3 and 4).
Populations of microorganisms were generally highest in 90-day old chestnuts at
the time of marketing (60 — 90 days after harvest) (Figures 3 and 4). In the case
of molds and MAB, microbial populations increased up to 8 and 8.3 logs CF Ulg
by day 60, and then decreased to 6.1 and 6.7 logs CF U/g after 120 days of
storage, respectively. During storage, mold (F (3,4) = 177.59, MSE = 6.72, p <
0.01) and MAB (F (3,4) = 206.44, MSE = 5.27 p < 0.01) populations from
chestnuts harvested directly from the tree were signiﬁcantly lower compared to
chestnuts harvested from the ground. On day 0 and beyond, mold populations on
the shell were signiﬁcantly higher (F (1,4) = 199.19, MSE =12.41 p < 0.01), than
the kernel (Figure 3). MAB populations on the shell were signiﬁcantly higher than
the population in the kernel (F (1 ,4) = 197.15, MSE =5.31 p < 0.01) (Figure 4).

Yeast populations did not reﬂect the same pattern in that the kernels of
chestnuts harvested from the tree were signiﬁcantly less (F (1 ,4) = 2.81, MSE
=1.50, ns), contaminated than those harvested from the ground (Figure 5). An
irregular growth rate and significant population increase (F (1 ,4) = 189.78, MSE
=17.29, p < 0.01) was observed in yeasts colonizing the shell. With these
populations on chestnuts collected from the orchard ground, signiﬁcantly
increasing (F (1 ,4) = 189.78, MSE =17.29, p < 0.01) to 8.5 logs CFU/g during 120
of storage. Meanwhile, yeast populations from the shell of chestnuts collected
directly from the tree, were more variable with a decline in population, from 4.1
logs CFU/g on day 30 down to 2.3 logs CFU/g 90 days later. Neveitheles's, after

120 days a signiﬁcant increase (F (1 ,4) = 189.78, MSE =17.29, p < 0.01) up to

63

6.7 logs CFU/g was observed. During storage, yeast populations on the shell was

signiﬁcantly higher (F (1 ,4) = 256.65, MSE =8.71 p < 0.01) compared to those in

the kernel. Populations of yeast on kernels collected from the tree or ground

remained relatively constant during storage (Figure 5).

 

 

 

a
9
Q l
o - a
_ {IAZ
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. Y I
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ab . -+ ........ 1C
. b ' - . - - ‘ .930
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_—" 0 30 thﬂﬂﬂw”*mgo' W~~ #126”
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.... Kemel-Tree

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Figure 3. Mean mold counts (log cfu g") on shells and In kernels of fresh chestnuts during
120 days of storage at 4° C. Data points followed by the same lower case letter within the same
day are not signiﬁcantly different at p = 0.05 (ANOVA with post-hoe Tukey multiple comparison of

means).

2Overall data point followed by the same capitalized letter within chestnut part-

harvesting method are not signiﬁcantly different at p = 0.05 (Repeated measurement design —
ANOVA with post-hoc Tukey multiple comparison of means). Error bars indicate standard

deviation.

64

 

 

 

 

a
a t. o Shell-Ground
‘ ‘ ... Shell-Tree
°° “ - KemekGround
Kemel-Tree
N a
+ A2
.. a
:6 b i
a ab 6 $\b
9;” 1 , , l/T,‘ i {B
8, 8/, ' '1'. ~_ ‘ I ,bi C
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v F ' ‘ ‘ ' '
.' c
9 l0
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l --- .___ __ _- ..
O 30 60 90 120
Storage (Days)

Figure 4. Mean MAB counts (log cfu g") on shells and In kernels of fresh chestnuts during

120 days of storage at 4° C. Data points followed by the same lower case letter within the same
day are not signiﬁcantly different at p = 0.05 (ANOVA with post-hoc Tukey multiple comparison of
Overall data point followed by the same capitalized letter within chestnut part-

means).

harvesting method are not signiﬁcantly different at p = 0.05 (Repeated measurement design -
ANOVA with post-hoc Tukey multiple comparison of means). Error bars indicate standard

deviation.

65

 

 

 

 

 

3.. 2 A Shell-Ground
. (A +— Shell-Tree
. l . ‘ -.-. Kernel-Ground
°° f ‘ ' ‘ Kemel-Tree
l I
a be I
o g A
1
as '3 I:
E a§__—————-- \9
09
:3: cl: c I;
N ?__,.---"'c§:-::-: ...... EL“... ----- 3P: ---------- EC
b ' b" c C
c.
To“ " * so _- 60 9.7“” mm“ ‘ W *
SIOFGQNDGYS)

Figure 5. Mean yeast counts (log cfu g") on shells and In kernels of fresh chestnuts during
120 days of storage at 4° C. 1Data points followed by the same lower case letter within the same
day are not signiﬁcantly different at p = 0.05 (ANOVA with post-hoe Tukey multiple comparison of

means). Overall data point followed by the same capitalized letter within chestnut part-
harvesting method are not signiﬁcantly different at p = 0.05 (Repeated measurement design —
ANOVA with post-hoc Tukey multiple comparison of means). Error bars indicate standard
deviation.

Farm inﬂuence on microbial populations on shell and in the kernel of

chestnuts

To determine the effect of different growing and harvesting conditions on
postharvest microbial populations of chestnuts on various farms in Michigan,
chestnuts were collected from seven different growers at the Chestnut Growers,
inc. on day of delivery. Populations on shell varied from farm to farm, with MAB,
molds and yeasts ranging from 4.55 to 8.44, 3.94 to 7.74, 4.75 to 8.28 logs

CFU/g respectively (Figure 6). A similar scenario was observed for the kernels

66

with MAB, molds and yeasts ranging from 2.34 to 7.63, 1.99 to 7.69, 2.68 to 7.24
logs CFU/g respectively (Figure 7). A signiﬁcant difference was observed on
molds (F (6,1) = 11.43, p < 0.01), MAB (F (6,1) = 9.85, p < 0.01) and yeasts (F
(6,1) = 27.82, p < 0.01) on the shell (Figure 6). And in the kernel (F (6,1) = 11.44,
p < 0.01), MAB (F (6,1) = 4.39, p = 0.01) and yeast (F (15.04) = 19.64, p < 0.01)
(Figure 7). Based on multivariate analysis of variance (MANOVA - ‘Wilks’ test), all
of the microbial populations (dependent variables) were signiﬁcantly different
from farm to farm, indicating a signiﬁcant difference between microbial
populations on shell (approx F (18,3) = 8.92, p < 0.01) (Figure 6) and kernel
(approx F (18,3) = 11.44, p < 0.01) (Figure 7). Chestnuts from farm C yielded
signiﬁcantly higher microbial population compared to those from farm D. With the

remaining farms being between farms C and D.

67

 

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Figure 8. Total MAB, molds and yeast counts (log cfu 9'1) from the shells of fresh
chestnuts after harvest from seven Michigan farms. 1Values followed by the same letter within

0.05 (ANOVA) (Tukey multiple comparison of

organisms are not Signiﬁcantly different at p =
means). Error bars indicate standard deviation.

68

 

 

 

 

 

 

 

 

 

 

 

 

 

 

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l 5 Yeast
o ‘ L. ...,.-__-. __ _ ”4
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A a c e F e

 

Growers

Figure 7. Total MAB, molds and yeast counts (log cfu g") from the kernels of fresh

chestnuts after harvest from seven Michigan farms. 1Values followed by the same letter within
organisms are not signiﬁcantly different at p = 0.05 (ANOVA) (Tukey multiple comparison of
means). Minimum detectable level (MDL) = Minimum possible count (0.5) x minimum dilution
factor x inoculated aliquot (100 pl). Error bars indicate standard deviation.

Farm inﬂuence on microbial quality of chestnuts

To determine the effect of different growing and harvesting conditions on
the severity of postharvest mold and kernel decay, chestnuts were collected from
the seven different growers when received at Chestnut Growers, Inc. On
receiving day (day 0) a signiﬁcant difference in mold severity (F (6,1) = 2.89, MSE
= 1.20, p = 0.03) and kernel decay incidence (F (6,1) = 2.61, MSE =0.11, p =

0.04), was observed between growers (Figure 8).

69

 

 

2:3 Shell-mold-severlty i
i I] Kernel-decay-incidence 1
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b
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1 p I - ..t ""
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Growers

Figure 8. Shell mold severity and kernel decay incidence from seven Michigan farms at

receiving day (Day 0). 1Values followed by the same letter within quality index are not
signiﬁcantly different at p = 0.05 (Repeated measurement design - ANOVA) (Tukey multiple
comparison of means). Error bars indicate standard deviation.

Overall, during 60 days of storage a signiﬁcant difference in mold severity
was observed on shells (F (6,2) = 5.03, MSE = 0.28, p < 0.01) but not in kernel
decay incidence (F (6,1) = 2.78, MSE = 0.04, ns), between growers (Figure 9).
Farms C and B had signiﬁcantly higher shell mold severity compared to the other

farms, with values of 2.8 and 1.5 logs CFU/g respectively.

70

4w “-..—.....- _._.._._ —.-—— H—._.s

 

 

 

 

 

 

 

 

 

W? 1 if: Shell-mold-severity '1
1 1 [I] Kernel-decay-incidence
1 -
(‘0 * l
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Growers

Figure 9. Shell mold severity and kernel decay incidence from seven Michigan farms,
during 60 day storage at 4°C. 1Values followed by the same letter within quality index are not

signiﬁcantly different at p = 0.05 (Repeated measurement design - ANOVA) (Tukey multiple
comparison of means). Error bars indicate standard deviation.

Discussion

Postharvest shell mold and kernel decay affects a signiﬁcant portion of
chestnuts worldwide and about 20 percent of the chestnuts produced in
Michigan. Results showed that fungi, bacteria and yeast signiﬁcantly increased
during storage. The extent of increase was dependent on the part of the chestnut
(shell or kernel) and the collecting method (tree or ground). Highest populations
were found on the shells of chestnuts collected from the ground. These results

are not surprising as the shell is continually exposed to the environment while the

71

kernel is protected by the shell and pellicle. Molds were the most noticeable
organisms associated with the shell and kernel decay of chestnuts during
refrigerated storage (4° C). Similar problems, related to mold decay also have
been reported in Asia, Europe, South America and North America (Jerimini et al.,
2006; Jian et al., 2002; Montealegre and Gonzalez, 1986; Miller, 2003; Paglietta
and Bonous, 1979; Rutter et al., 1990; Vettraino et al., 2005; Vossen, 2000;
Washington, 1997). The fungal ﬂora responsible for this infestation is diverse and
appears to be strongly inﬂuenced by preharvest, harvest and storage conditions.
In the traditional method of harvest, chestnuts that have fallen to the ground are
collected by the workers or mechanical pickers (Anagnostakis, 2003). Attempts
have been made to collect chestnuts in nets either on the ground or suspended
above the ground, in italy and France mainly to ease chestnut collection (Breisch,
1993; Bonous and Raccolta, 2002) and reduce molding, especially by Sclerotinia
pseudotuberosa (Breisch, 1993). However, recent study indicated that
suspended nets did not reduce chestnut mold after harvest or during storage
(Sieber et al., 2007). Regardless, in this study, chestnuts harvested from the
orchard ﬂoor were more heavily contaminated during storage, than those
harvested directly from the tree.

Microbial populations are much higher on chestnut shells compared to the
kernels. In general, only two fungal genera caused obvious mold symptoms with
chestnuts shells after harvest and during storage. in order of importance, these
were Penicillium spp. (P. expansum, P. griseofulvum, P. chysogenum) and

Coniophora puteana. However, at least ﬁve different genera of ﬁlamentous fungi

72

were associated with infected chestnut kernels. These microorganisms which
included Penicillium spp. (P. griseofulvum, P. expansum, P. chysogenum),
Acrospaeria mirabilis, Botryosphaeria ribis, Sclerotinia sclerotiorum, Botryotinia
fuckeliana (Anamorph Botrytis cinerea) and Gibberella sp. (Anamorph Fusarium
sp.) were generally undetectable and caused the largest amount of loss. Other
organisms might be associated with chestnut shell mold and kernel decay in
Michigan, but during this two-season study, only these species were repeatedly
associated with postharvest shell mold and kernel decay. Similar ﬁndings have
been reported for chestnuts harvested in France, Italy, Australia, Chile, United
States and other countries (Ellis and Ellis, 1985; Jerimini, et al., 2006;
Montealegre and Gonzalez, 1986; Paglietta and Bonous, 1979; Ridé and Gudin,
1960; Vettraino et al., 2005 Washington, et al. 1997; Wright, 1960).

This was the ﬁrst survey to associate Coniophora puteana, Botryosphaen'a
n'bis, and Sclerotinia sclerotiorum with chestnut mold or kernel decay.
Furthermore, S. sclerotiorum, which was found infrequently in our survey, has
never been previously reported on chestnuts, but symptoms and pathogenesis
may be similar to that of Sclerotinia pseudotuberosa, mold responsible for
chestnut black rot, which is one of the most important postharvest diseases of
acorns (Quercus spp.) in Europe (Vettraino, et al., 2005; Washington et al.,
1997).

Findings concerning differences between mold severity in shell and
microbial populations on chestnuts among farms, demonstrated that location,
preharvest and harvesting conditions played an important role in quality and

decay of chestnuts (Kader, 2002; Sieber, et al. 2007; Willis, et al., 2007).

73

Variation in microbial populations among farms, increases in populations during
storage and variability of populations within the same ﬁeld during storage,
suggest multiple preharvest sources of contamination including soil, feces,
irrigation water, water used to apply fungicides and insecticides, insects,
inadequately composted manure, wild animals, and human handlers (Beuchat,
1996). Therefore, complementary studies on the efﬁcacy of preharvest
management, preharvest fungicide applications, and path of produce
contamination, as well as harvesting methods and storage conditions must be
considered. Furthermore, the interactions between climate and epidemiology of
the fungal ﬂora in the ﬁeld also need to be better understood. Currently, all these
factors must be taken into consideration and conﬁrm the need to control the
organisms prior to storage to avoid or reduce postharvest mold and decay during
storage. These efforts must be in concert with a plan to efﬁciently identify and

manage affected products during harvest and storage.

74

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American Public Health Association (APHA). (2001). Compendium of methods for
the microbiological examination of foods (4th. Edition ed.). Washington,
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Anagnostakis, S. L. (1999). Cultivars of chestnuts. Annu. Rpt. Northern Nut
Grower Assn. (90), 16-31.

Anagnostakis, S. L., and Devin, P. (1999). Nutrients in chestnuts. Annu. Rpt.
Northern Nut Grower Assn. (90), 36-40.

Anagnostakis, S. L., Gordon, R, and Hebard, F. V. (1998). Identiﬁcation of
chestnut trees. Annu. Rpt. Northern Nut Grower Assn. (89), 1-4.

Beuchat, L. R. (1996). Pathogenic microorganisms associated with fresh
produce. Journal of Food Protection, 59, 204-216.

Biomhoff, R., Carisen, M. H., Andersen, L. F., and Jacobs, D. R. (2006). Health
beneﬁts of nuts: potential role of antioxidants. British Journal of Nutrition,
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Blackwell, R. (2006). Chestnut Farms, Inc. Business plan. Chestnut Farms, Inc.
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79

CHAPTER 3

EFFICACY OF POSTHARVEST TREATMENTS FOR REDUCTION OF MOLD
AND DECAY IN FRESH CHESTNUTS

Abstract

In Michigan where chestnut (Castanea spp.) cultivation is a pioneering
industry, postharvest molds are responsible for signiﬁcant economic losses.
Molds contaminate chestnut before and at time of harvest and are exacerbated
during transportation and storage. Consequently, various chemical sanitizers for
fresh produce were evaluated for their effect on management of postharvest
chestnut shell mold, kernel decay and weight loss during storage. Treatments
were evaluated in chestnuts randomly collected after harvest from seven
Michigan growers. Each treatment was repeated four times with 2.3 kg chestnuts
per replication. Per replication a sub-group of 10 chestnuts was randomly picked
and evaluated for microbiological quality (severity of shell mold and incidence of
kernel decay) and weight loss. Overall, 2,700-ppm hydrogen peroxide + 200-pp,
peracetic acid and 0.15-ppm triﬂoxystrobin signiﬁcantly reduced both mold
severity on the shell and the incidence of internal kernel decay compared to the
other products. Other products and combinations of products may also play an
important role in reducing the incidence of decayed kernels, even though they did
not strongly reduce mold severity. Among these, were 0.92-ppm ozone solution,
7.5-mg chlorine dioxide/g of chestnuts, 100-ppm peracetic acid, 0.2-M sodium

metabisulﬁte, and 1,350-ppm hydrogen peroxide + 100-ppm peracetic acid in

80

combination with 1,980-ppm caprylic acid, 1,333-ppm glycolic acid and 20-ppm
capric acid. No product signiﬁcantly reduced weight loss during storage. Based
on these ﬁndings 2,700-ppm hydrogen peroxide + 200-ppm peracetic acid and
0.92-ppm ozone solution are best suited to protect against postharvest chestnut
mold and kernel decay when combined with good agricultural practices and

postharvest management.

Introduction

Postharvest decay of edible chestnuts (Castanea spp.) reduces nut quality
and can lead to severe economic losses (Jerimini, et al., 2006; McCarter et al.,
1980; Narayanasamy, 2006). A diverse range of ﬁlamentous fungi including
Penicillium sp., Aspergillus sp., Fusarium sp., Phomopsis castanea, Acrospeira
mirabilis, and Sclerotinia pseudotuberosa (syn. Ciboria batschiana, S.
batschiana; anamorphic from Rhacodiella castanea, syn. Myrioconium castanea)
have been most commonly responsible for postharvest decay of the shell and
kernel in France, Italy, Australia, Chile, United States and elsewhere (Ellis and
Ellis, 1985; Jerimini et al., 2006; Montealegre, J. and Gonzalez, 8. 1986;
Paglietta and Bonous, 1979; Ride, M. and Gudin, C. 1960; Sieber, Jermini, and
Conedera, 2007; Vettraino et al., 2005 Washington et al. 1997; Wright, 1960). In
Michigan, postharvest decay has been identiﬁed as one of the major problems
that negatively impacts fresh chestnuts, accounting for up to 25 percent of losses
after harvest. Recently, several fungal species, have been isolated from fresh
healthy Michigan chestnuts and chestnuts experiencing postharvest shell mold

and kernel decay, including Penicillium spp. (P. griseofulvum, P. expansum, P.

81

chysogenum), Acrospaeria mirabilis, Botryosphaen'a n'bis, Sclerotinia
sclerotiorum, Botryotinia fuckeliana (Anamorph Botrytis cinerea), Gibberella sp.
(Anamorph Fusarium sp.), and Coniphora puteana (Chapter 2, this thesis).
Various chemical and non-chemical treatments have been previously

evaluated for their impact on the quality of fresh chestnuts during storage. Among
the treatments tested, 95 % alcohol, 1 % copper sulfate, 0.4% calcium chloride,
boric acid, formaldehyde, benzoic acid, sulfur dust and others were unable to
prevent storage molds (Hammer, 1949; Tan et al., 2007). Furthermore, certain
treatments (boric acid and formaldehyde) compromised the chestnut quality by
hardening the kernel (Hammer, 1949). Other chemical treatments potentially are
able to minimize postharvest decay, including 30-ppm iodine, 250-ppm
carbendazim, 100 to 200—ppm sodium hypochlorite, 100-ppm peracetic acid
(Morris, 2006; Panagou, Vekiari, and Mallidis, 2005), 50-ppm natamycin, 1 %
sorbic acid, 1 % propionic acid, hot water dip at 90° C for 10 min (Panagou,
Vekiari, and Mallidis, 2005) and immersion in fungicides including 2,6-dichloro-4-
nitroaniline (Botran) (Wells, 1980). Immersion in 15° C water for 15 min, 4548 °
C water for 15 min and 52° C water for 5, 15, 30 or 60 min signiﬁcantly reduced
both the percentage of infected nuts during storage and the percentage of
chestnuts exhibiting insect damage from the presence of larvae of Cydia
splendana, Curculio elephas and Curculio sayi. (Jerimini, et al., 2006; Sieber,
Jermini, and Conedera, 2007; Wells, 1980).

While various strategies can reduce both microorganisms and insects,

some of these treatments, such as immersion in 52° C water for 60 min, will

82

decrease soluble sugars and increasing starch during storage, negatively
affecting chestnut quality (Wells, 1980). Improved food processing and
management practices can be used to enhance the safety, storage and shelf life
of various commodities. These strategies primarily involve various sanitizers such
as hydrogen peroxide, ozone, chlorine dioxide (gas and solution), hypochlorite,
organic acids, natamycin, along with heat (Beuchat 1998; Block 1991; Brackett
1999; Crowe et al., 2005; Palou et al., 2002; Panagou et al., 2005; Perrera and
Karunaratne 2001; Pierre et al., 2006, Sapers 2003; Suslow 2002; Wade et al.,
2003). However several salts, oils and biocontrol agents have also been
evaluated for reducing decay (Feng and Zheng, 2006, Kader, 2002; Mari et al.,
2003; Mills et al., 2004; Porat et al., 2002; Sapers, 2003; Willis et al., 2007).

With the objective of understanding the effectiveness of postharvest
treatment on microbial quality of Michigan fresh chestnuts, including the reduction
of shell mold, kernel decay and weight loss during storage, sixteen different

sanitizers were evaluated.
Material and Methods

Samples

A total of 454 kg Chestnuts (cv. ‘Colossa|’) were obtained immediately
after harvest, from seven commercial farms in Michigan and were used after 2
days of storage at 4 °C (MSU experimental station, Clarksville, MI, USA). All
chestnuts were completely mixed, randomized and placed in pure water (~ 5,000
It) where the majority of decayed, empty or damaged chestnuts were eliminated

by their proclivity to ﬂoat, as healthy chestnuts tend to sink.

83

Treatments and storage

Sixteen different treatments were tested are described in Table 3, with

each treatment replicated four times using 2.3 kg of chestnuts.

Table 3. Postharvest treatments used in fresh chestnuts

 

 

 

 

 

 

 

 

 

 

Exposure
Commercial name Active Ingredient Concentration time (min)
Scholar Fungicide 50% F ludioxonil 0.03 ppm 3
(Syngenta Crop
Protection, Inc.
Greensboro, NC, USA)
Flint Fungicide (Bayer 50% Triﬂoxystrobin 0.15 ppm 3
CropScience Int.)
CAP-06m (Bjosafe 39.6% Caprylic acid, 26.67 3,960 ppm 3
Systems, Glastonbury, emsorb6915, 14% glycolic caprylic acid,
CT, USA) acid, 13.3 light mineral oil and 2,667 glycolic
0.4% capric acid acid, 40 ppm
capric acid
Storoxm (Biosafe 27% Hydrogen dioxide + 2% 2700 ppm 3
Systems, Glastonbury, Peracetic acid hydrogen
CT, USA) dioxide + 200
ppm peracetic
acid
smroxTM + CAF-06 TM 27% Hydrogen dioxide + 2% 1350 ppm 3
(Biosafe Systems, Peracetic acid + 39.6% hydrogen
Glastonbury, CT, USA) Caprylic acid, 26.67 dioxide 1' 100
emsorb6915, 14% glycolic ppm 2 peracetic
acid, 13.3 light mineral oil and acid + 1,980
0.4% capric acid ppm caprylic
acid, 1,333
glycolic acid, 20
ppm capric acid
Agri-Cide (Life Science 18.25 - 21.75% Copper 1 ppm 3
Group, Inc. Monticello, sulfate pentahydrate
IN, USA)
Chlorine dioxide gas (3102 75 mg of 0102/9 120
(ICA TriNova, LLC chestnut
Forest Park, GA)
Chlorine dioxide solution c|02 + H20 10 ppm 3
(lCA TriNova, LLC
Forest Park, GA)
Ozone solution (Aqua Air 03 + H20 0.92 ppm 3

Technologies, Inc.,
Bloomﬁeld, ML USA)

84

Table 1. (cont’d)

 

 

 

 

Commercial name Active ingredient Concentration Exposure
time (min)

Sodium metabisulﬁte N323205 0.2 M 3
(Sigma-Aldrichm, St.
Louis, MO, USA)
Aluminum acetate (CH3C02)2 AIOH 0.2 M _ 3
(Sigma-Aldrich”, St.
Louis, MO, USA)
Bio-Save® 11 LP Pseudomonas syringae 1.65 g/L of 3

Biological fungicide
(Jet harvest solutions,
Longwood, FL, USA)

Strain ESC-11

water

 

RadianceTM (CHzO,
Olympia, WA, USA)

Shellac-based wax

Concentrated 45

 

Brilliance 3TM - CH20,
Olympia, WA, USA

Caranauba-based wax

Concentrated 45

 

Peracetic acid
(Len ntech 7”,

Rotterdamseweg,
Netherlands)

Peracetic acid

100 ppm 3

 

Chlorine solution
(Champion packaging,
Inc. Woodridge, IL, USA)

6% Sodium hypochlorite

1000 ppm of 3
sodium
hypochlorite

For each treatment a sub-group of 10 chestnuts was randomly picked and

then tightly wrapped in a manually cut mesh bag (Figure 9-a). This sub-sample

was then placed in the middle of remaining chestnuts containing 2.5 kg of

chestnuts, thus creating a double-sac where a larger group of chestnuts from the

same treatment surrounded the small sub-sample of 10 chestnuts, thereby

protecting them from contact with untreated chestnuts and chestnuts receiving

different treatments (Figure 9-b) (Ryser, 2007). All treatments solutions, except

chlorine dioxide gas, ozone and waxes were thoroughly mixed, to achieve the

desired concentration in 30 L of water. Experiment was performed once

consisting in four samples per treatment, which were dipped at once in each

solution for 3 min (exposure time) (Figure 9-c).

85

Chestnuts were exposed to 7.5 mg of ClOz gas lg of chestnuts inside a 20
L sealed bucket for 2 h (exposure time). Chlorine dioxide gas was generated in-
situ using a 24 9 commercial ClOz sachet (ICA TriNova, LLC Forest Park, GA)
and circulated inside the sealed bucket using three 12 VDC/0.07-0.11A cooling
fans (40x40x10 mm-case-fan) (Model MW-410H12C, AAVID, Bologna, Italy) that
were attached to the bottom and opposite sides of the bucket (Figure 9-f). Ozone
was generated using a commercial ozone generator (ozone solution, Aqua Air
Technologies, Inc., Bloomﬁeld, MI, USA) equipped with an oxygen concentrator
(model CD 10/AD Corona Discharge ozone generator system, Clear Water
Tech., Inc., San Luis Obispo, CA, USA). The gas was delivered through an inlet
line directly into the water, which was constantly circulated through a 30 L
container. Water containing up to 1-ppm ozone was produced within 20 min. The
concentration of ozone in water used to treat chestnuts was constantly monitored
by an electrochemical method (Protti, 2001) using a portable dissolved ozone
meter (model OZ-21 P, DKK—TOA Corporation, Takadanobaba, Shinjuku-ku,
Tokyo, Japan).

Chestnuts were spray-waxed on clean Teletype paper (PM Company,
Cincinnati, OH, USA) (Figure 9-e) using a manual pump spraying system model
Jr. pump-up (JaniSan, Layton, UT, USA). After air-drying for 45 min, excess wax
was manually removed with a clean polypropylene scrub brush (Fuller, Great
Bend, Kansas, USA).

All treated samples were then stored submerged in one full 453.6 kg high-

density polyethylene vented bin (MACX, Grand Rapids, MI, USA) of untreated

86

chestnuts which was held at 4 °C using a completely randomized design into four
vertical strata (1 repetition per strata) (Figure 9-d) (MSU experimental station,
Clarksville, MI, USA). Temperature (T) and relative humidity (RH) were
continuously monitored in the 4 strata, using a data logger (Model: HOBO Micro
station data logger - H21-002 with 4-Temp/RH sensor — S-THB-M002,

MicroDAQ, Contoocook, NH, USA).

 

 

A . O ... ‘ “r I. 0‘ in 3‘ .
=igure 10. Treatment of fresh chestnuts before storage. a) sue-sample of 10 chestnuts, b)
double-sac storage method, c) liquid-solution treatment application, (I) Chlorine dioxide

gas system used to treat chestnuts, a) wax application, f) treatment storage. Image In this
thesis Is presented in color.

 

 

 

 

 

 

Microbial mold severity, kernel incidence and weight loss analysis
Microbiological quality (severity of shell mold and incidence of kernel

decay) and weight loss from chestnuts in the sub-sample of all 16 treatments was

87

evaluated after 60 days storage. Remaining chestnuts in treated samples were
discarded. Shell mold severity was calculated as the mean number of a 4-point
visual qualitative measurement of mold presence, where 0 = no mold, 1 = 1-25 %
mold, 2 = 26-50 % mold, 3 = 51-75 % mold and 4 = 76-100 % mold. Incidence of
decayed kernels was calculated as the mean number of chestnuts exhibiting

decay. Weight loss was the difference from the in weight and after storage.

Statistical analysis

The mold severity, kernel decay and weight loss data collected after 690
days of storage were analyzed using one-factor analysis of variance (ANOVA.
Signiﬁcance between means was determined using the Tukey post-hoc multiple
comparisons of means test at the 95% family-wise conﬁdence level (p=0.05).
Calculations were performed using the “R: A language and environment for
statistical computing” statistical package (R Development Core Team, 2007) (Ott

and Longnecker, 2001).

Results

A statistically signiﬁcant difference was observed for shell mold severity (F
(15,1) = 0.89, p < 0.01) and incidence of decay in kernels (F (15,1) = 4.88, p <
0.01) (Table 4), conﬁrming that at least one treatment was statistically lower or
higher in comparison with the other treatments. The most efﬁcacious treatments
for inhibiting shell mold included 3 minutes immersion time in 2,700-ppm

hydrogen dioxide + 200-ppm peracetic acid, 0.15-ppm triﬂoxystrobin and 1,350-

88

ppm hydrogen dioxide + 100-ppm peracetic acid mixed with 1,980-ppm caprylic
acid, 1,333-ppm glycolic acid and 20-ppm capric acid.

Table 4.Effect of fresh chestnut treatment application on shell mold1, kernel decay2 and
weigh loss3 stored for 60 days in control environment4

Mold severity on Incidence of

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

Treatment shell“ decay in kernel Weight loss (9)
2700 ppm hydrogen dioxide 0.17 t 0.12 a 1.25 i 0.95 a 0.38 :I: 0.08 a
+ 200 ppm peracetic acid
0.15 ppm Triﬂoxystrobin 0.20 d: 0.27 a 1.50 :t 1.33 a 0.40 i 0.08 a
1350 ppm hydrogen dioxide 0.25 1: 0.19 a 3.50 1: 0.57 ab 0.22 :t 0.19 a
+ 100 ppm 2 peracetic acid
+ 1,980 ppm caprylic acid,
1,333 glycolic acid, 20 ppm
capric acid
0.03 ppm Fludioxonil 0.27 1 0.15 ab 1.25 1 0.95 a 0.30 t 0.13 a
3,960 ppm caprylic acid, 0.40 :l: 0.21 b 2.75 i 1.50 ab 0.27 :l: 0.09 a
2,667 glycolic acid, 40 ppm
capric acid
10ppm (3102 0.40 t 0.31 b 2.75 :I: 0.95 ab 0.27 :I: 0.17 a
0.2 M Sodium metabisulﬁte 0.48 1 0.12 c 2.25 :l: 1.50 a 0.25 :t 0.05 a
1000 ppm Sodium 0.58 t 0.09 c 3.75 :I: 0.96 ab 0.27 :I: 0.12 a
hypochlorite
0.2 M Aluminum acetate 0.63 t 0.17 cd 3.25 i 2.06 ab 0.35 :t 0.13 a
1 ppm Agri-Cide 0.85 t 0.26 d 4.25 :t 1.50 ab 0.30 :l: 0.14 a
2 L-concentrated Brilliance 0.98 :t 0.15 d 3.00 :t 1.41 ab 0.27 :I: 0.15 a
wax
0.92 ppm Ozone 1.12 :t 0.17 d 0.50 :l: 0.58 a 0.35 :t 0.10 a
2 L-concentrated Radiance 1.12 t 0.52 d 3.25 :t 1.50 ab 0.17 :t 0.15 a
wax
7.5 mg CIOz 99319 chestnut 1.23 :t 0.26 d 1.25 :l: 0.50 a 0.20 t 0.16 a
16.5 g Bio-save 11 LP/L 1.28 :l: 0.15 d 4.50 :t 0.58 b 0.27 t 0.05 a
water
100 ppm PAA 1.40 :t 0.36 d 2.00 :t 0.82 a 0.27 :r: 0.19 a
Non-treated Control 1.68 i 0.12 d 5.50 :l: 1.73 b 0.2 :t 0.11 a

 

 

 

1Shell mold severity was calculated as the mean number of a 4-point visual qualitative
measurement of mold presence, where 0 = no mold, 1 = 1-25 % mold, 2 = 26-50 % mold, 3 = 51-
75 % mold and 4 = 76-100 % mold
2Incidence of decayed kernels was calculated as the mean number of chestnuts showing
symptoms of decay (n=10).

eight loss was calculated as the mean of total weight before storage minus weight after
storage.
4Treatments stored submerged in one full 453.6 kg high-density polyethylene vented bin of
untreated chestnuts held at 4 °C using a completely randomized design into four vertical strata (1
repetition per strata) (mean T/R.H. = 7.2°C/96.5%).
5Numbers followed by the same letter within the column are not signiﬁcantly different at p = 0.05
(Tukey multiple comparison of means).

89

Treatments that signiﬁcantly reduced the number of decayed kernels in
comparison with the non-treated control and other treatments were 0.92-ppm
ozone, 7.5-mg ClOz gas/g chestnut, 0.03-ppm ﬂudioxonil, 2,700—ppm hydrogen
dioxide + 200-ppm peracetic acid, 0.15-ppm triﬂoxystrobin and 100-ppm
peracetic acid. Of the evaluated treatments, none signiﬁcantly reduced weight

loss during storage (F (15,1) = 0.02, ns).

Discussion

Currently, the Michigan chestnut industry immerses harvested chestnuts in
a 100-200 ppm chlorinated water bath to reduce contaminants and microbial
population before storage. However, preliminary research indicated that the
efﬁcacy of this compound is not only limited, but may also have unpredictable
side effects (Cena, 1998; Nguyen-rhe and Carlin, 1994). In recent years,
important health and environmental concerns have arisen over the continued use
of chlorine due to its reactivity with some food constituents to form toxic by-
products (Cena, 1998; Graham, 1997). Thus, with the actual regulations for
chemical use and possibility of future regulatory constraints on the use of chlorine
as a sanitizer, alternative treatments are being sought to improve the
microbiological quality and safety of agricultural commodities. Other laboratories
including Beuchat (1998), Block (1991), Brackett (1999), Crowe et al. (2005),
Palou et al. (2002), Panagou et al., (2005), Perrera and Karunaratne (2001),
Pierre et al., (2006), Sapers (2003), Suslow (2002), and Wade et al., (2003),
have evaluated other potentially safer sanitizers such as hydrogen peroxide,

diverse acids, ozone, chlorine dioxide (gas and solution), hypochlorite, organic

90

acids and natamycin in addition to heat for enhancing the safety and shelf of a
wide range of commodities. Several salts, oils and biocontrol agents that can
reduce decay are also being used and evaluated (Feng and Zheng, 2006, Kader,
2002; Mari et al., 2003; Mills et al., 2004; Porat et al., 2002; Sapers, 2003; Willis
et a., 2007).

The present work was conducted to evaluate the efﬁcacy of 16 postharvest
treatments, with the objective of improving microbiological quality of fresh
chestnuts. Hydrogen dioxide signiﬁcantly reduced both mold severity in shell and
incidence of kernel decay. Hydrogen dioxide has been used at concentrations of
1 to 10%, to reduce microbial populations and extend the shelf life many fruits
and vegetables (Crowe, Bushway, and Bushway, 2005; Sapers, 2003). This
compound is approved by the Food and Drug Administration (FDA) and it is
generally recognized as safe (GRAS), mainly because it produces no residues
since it is reduced to water and oxygen (Sapers, 2003). Although treatments
containing high concentrations of hydrogen dioxide may be beneﬁcial in
improving microbial quality and extending shelf life, use of this treatment may be
limited for lettuce, blueberries and other commodities due to excessive surface
oxidation, resulting in product damage (Aharoni, Copel, and Fallick, 1994; Block,
1991; Simmons, 1997; Ukuku and Sapers, 2001; Ukuku, Pilizota, and Sapers,
2001). Although hydrogen dioxide in combination with caprylic acid, glycolic acid
and capric acid also signiﬁcantly reduced mold shell severity, no reduction was
seen in reducing kemei decay. Similar results have been obtained for blueberries
and in green salads, where hydrogen dioxide in combination with citric acid

suppressed the growth of microbial populations and extended the shelf life of

91

both commodities (Crowe, Bushway, and Bushway, 2005; Sapers, 2003).
Furthermore, there is no evidence indicating that fresh chestnuts were negatively
affected by any of the previously mentioned products.

This study demonstrated that certain sanitizers might also play an
important role in reducing the incidence of decayed kernels, even though they
were not as effective in reducing mold severity. Among these were 0.92-ppm
ozone solution, 7.5-mg of chlorine dioxide gas/g of chestnut, 100—ppm peracetic
acid, and 0.2-M sodium metabisulﬁte. Ozone, chlorine dioxide gas and peracetic
acid are used in several sanitizing processes. They are all FDA approved and
have played important roles in reducing spoilage organisms, postharvest
pathogens and foodbome pathogens on ready-to-eat produce, and other diverse
horticultural commodities.

In general, the previously mentioned sanitizers more potent than chlorine
solutions, in FDA permitted concentrations (Gurol and Akata, 1996; Mari et al.,
1999; Morris, 2006; Palou et al., 2002; Pierre et al., 2006; Panagou et al., 2005;
Sapers, 2003; Roberts and Reymond, 1999; Sapers, 2003; Sarig et al., 1980;
Sharma et al. 2003; Suslow, 2002; Wade et al., 2003; Wu and Kim, 2007; Zoffoli
et al., 2005). Sodium metabisulﬁte, with relatively low mammalian toxicity and a
broad-spectrum mode of action, was also effective in reducing kernel decay. It is
widely used as a multi-functional antimicrobial agent and is a common
preservative found in relishes, fresh and dried fruits and vegetables, tomato
paste, mincemeat and potatoes (Mills et al., 2004; Porat et al., 2002).

Chemical treatments, including a broad range of fungicides such as

mancozeb, thiabendasole, ﬂudoxonil and triﬂoxystrobin can reduce decay in

92

different agricultural commodities (Gamage et al., 1997; Kader, 2002; Kicast,
1995; Narayanasamy, 2006; Sapers, 2003; Saroj, et al., 2006; Smith and Pillai,
2004). However, governmental regulations involving health and environmental
issues impair their use. This study showed that triﬁoxystrobin signiﬁcantly
reduced shell mold severity and the incidence of kernel decay. Fludioxonil
signiﬁcantly reduced the incidence of kemei decay, compared with the control
and other treatments, but was not among the best in reducing shell mold, when
compared with other products.

Wax coatings are used to the prolong shelf life of produce by decreasing
respiration rates, delaying fruit ripening, reducing shrinkage, reducing weight
loss, improving product appearance and providing a carrier for postharvest
treatment applications including the application of growth regulators. Edible
coatings have been tried on various fruits such as mandarins (Bayindirly,
Sumnu, and Kamadar, 1995), citrus fruits (Hagenmaier and Baker, 1993), pears
(Sumnu and Bayindirly, 1994), bananas (Alzaemey, Fallana, and Thomson,
1989) and apples (Chai, Ott, and Cash, 1991). This study demonstrated that
none of the postharvest treatments, including the two evaluated waxes
(caranauba- and shellac- base waxes) signiﬁcantly reduced weight loss of
chestnuts during storage. However, future studies may conﬁrm that the
appearance of the chestnuts might be improved, a factor that was observed, but
not measured in this study.

This study identiﬁed several compounds that may inhibit the growth of
microorganisms responsible for postharvest mold and decay of chestnuts. Among

these, was hydrogen dioxide, a product that could signiﬁcantly control

93

postharvest mold and decay of chestnuts. Other compounds with the potential to
inhibit internal decay include ozone, chlorine dioxide gas, peracetic acid and
sodium metabisulﬁte. The prospect of commercially using these compounds as
postharvest treatments needs to be further examined for potential advantages
over conventional chestnut production practices. Efﬁcient application and
monitoring will need to be developed and examined for these new microbial
reduction strategies to maximize product coverage and efﬁcacy. Most
compounds presently used in the food processing and agriculture industry are
generally regarded as safe for human consumption at the appropriate
concentration; hence hydrogen dioxide could be most easily adapted by chestnut

producers to reduce postharvest mold and decay.

94

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101

CHAPTER 4

MICROBIAL CONTAMINATION IN PEELED CHESTNUTS AND THE EFFICACY
OF POST-PROCESSING TREATMENTS FOR MICROBIAL SPOILAGE
MANAGEMENT

Abstract

Peeled chestnuts represent a growing value-added commodity worldwide.
Unlike most nuts, peeled chestnuts must be dehydrated or immediately frozen for
later use. Concerns regarding the shelf life and spoilage of peeled chestnuts after
thawing prompted a 2006-2007 survey in which chestnuts were quantitatively
examined for various microbial contaminants after harvest as well as during and
after peeling. Chestnuts (cv. ‘Colossal’) were randomly collected after harvest
from at least seven Michigan farms, stored at 4 °C for 2 weeks, and peeled using
a commercial brulage peeler. Microbial surveys indicated that average mesophilic
aerobic bacteria (MAB), yeast and molds populations in chestnuts were 2.70,
2.74 and 2.51 at post harvest; 3.46, 3.27 and 2.40 during peeling; and 5.39, 3.09
and <1.70 log CFU/g after peeling, respectively. Overall, microbial populations in
the water and environmental samples increased from < 1.7 to 5.46 log CFU/ml
and <17 to 5.91 CFU/10 cmz, respectively. During processing, the skin separator
(brushes) and a sorting belt were identiﬁed as key points for contamination.
Further studies indicated that two bacteria Rahnella sp., and Curtobacterium sp.
and the yeast Candida sp. were the primary causes of spoilage. Trials were
conducted to evaluate the efﬁcacy of six sanitizer treatments, as well as X-ray
irradiation and heat against these organisms on artiﬁcially inoculated chestnuts,

during 18 days of storage at 4 °C. Overall, X-ray irradiation, 3 min immersion in

102

92 ppm hydrogen dioxide and 65 °C hot water were most effective in reducing

MAB and yeast, compared to the other treatments.

Introduction

As an alternative to fresh chestnut sales, peeled chestnuts are processed
and sold to diversify the Michigan chestnut industry. Demand for products such
as peeled chestnuts, considered to be a fresh processed (cut) product, is
increasing as a result of consumer desire for healthy, fresh, easy to use and
appetizing foods (IFPA, 2006; Kader, 2002). In Michigan, fresh chestnuts are
commercially peeled with a commercial peeling system made in Italy and
purchased and imported in 2001 (Boema; Neive, Italy) (Appendix B). After
peeling, the chestnuts are typically vacuum-packed and stored frozen (Guyer et
al., 2003). Other mechanical methods of peeling, for example, air-impingement
de-shelling technologies, are also available but are not used in Michigan (Gao et
aL,2008)

Removing the shell and pellicle, which are considered to be a weak, but
still natural physical barrier that protects the kemei, facilitates water loss and
contamination of the kernel with pathogens and opportunistic organisms
(Cantwell, 1995; Mencarelli, 2001), potentially affecting their ﬁnal quality and
safety (Field, Jordan, and Osbourn, 2006).

Within ten to ﬁfteen days after thawing (~12 days), sticky, yellow ooze can
develop on the surface of thawed peeled chestnuts, affecting their ﬁnal quality
and acceptability. Similar problems have never been reported in peeled

chestnuts, but presence of slime or ooze is also common in processed fruits,

103

vegetables, juices, ready-to-use salads and meats (Brightwell et al., 2007;
Guerzoni et al., 1996; James, Martin, and David, 2005; N9, 2007; Ragaert et al.,
2006; Tournas, 2005).

Growth of spoilage organisms is usually accompanied by the accumulation
of metabolites, such as ethanol, lactic acid, and ethyl acetate, among others. The
organoleptic changes due to such metabolic activity are associated with the
enzymatic oxidation of various compounds leaking from the injured tissues.
Spoilage is detectable by sensory and microbiological methods when the
microbial population attains a certain microbial level. This population level is
dependent on the microbial species in question and the ingredient characteristics.
In extreme cases, shelf life of commercial products may not exceed ﬁve days
under conditions of refrigeration temperature (4 °C) (Brightwell et al., 2007;
Guerzoni et al., 1996; James et al., 2005; N9, 2007; Ragaert et al., 2006;
Tournas, 2005). Zagory (1998) and Tournas (2005) indicated that after harvest, a
wide range of economical important vegetables and speciﬁcally fresh-cut
products, are often spoiled by a wide variety of microorganisms including many
bacterial (Curtobacterium, Rahnella, Erwinia, Pseudomonas spp., among others)
and several fungal species. Spoilage signiﬁcantly increases when the product is
injured or the skin has been damaged or removed. Packaged sliced onions,
shredded mixed lettuce and other commodities stored under air or modiﬁed
atmosphere suffer from extreme colonization of diverse spoilage microorganisms,
including bacteria, molds and several yeasts (Pichia fennentans, Cryptococcus
Iaurentii and Candida spp., among others) (Liu and Li, 2006; Ragaert et al.,

2006). All these usually lead to undesirable quality, economic loss and conﬁrm

104

the importance of microbial growth during production, harvest, processing, and
during storage (Tournas, 2005; Zagory, 1998).

Increasing attention has been focused on the microbial safety of
processed fruits and vegetables, mainly on intervention methods to kill or remove
human pathogens and spoilage microorganisms in fresh produce (Perish, et al.,
2003; Sapers, 2003). In the majority of cases, sanitizing agents are added to
processing water to reduce microbial populations and prevent cross-
contamination of the product. Of these, chlorine has been used for several
decades and is still the most widely used sanitizer in the food industry (Baur et
al., 2004; IAFP, 2002). However, chlorine does not always reduce microbial
populations, including foodborne pathogens and may be harmful due to the
formation of toxic chlorine byproducts (Foley et al., 2004; Richardson, et al.,
2000; Vitro et al., 2005). New technologies, i.e., food irradiation, heat and others,
which have been reported to achieve greater than a 5-Iog reduction of pathogens
and other microorganisms (pasteurization treatments), have the potential to
greatly improve the microbiological quality and safety of produce (Niemira, 2003).

The objectives of this study were to determine which spoilage organisms
negatively impact peeled chestnuts before, during and after processing and how
they were becoming established on the processed product. In addition, the
efﬁciency of various postprocessing treatments to reduce microbiological ~

populations and prolong the shelf life of peeled chestnuts was also assessed.

105

Material and Methods

Chestnut samples

During the 2006 and 2007 growing seasons, chestnut samples (cv.
‘Colossal’) from at least seven Michigan farms were assessed for microbial
populations at two points before processing and ﬁve points during commercial
processing (Boema; Neive, Italy), at the Michigan State University Rogers
Reserve, Jackson. MI. Triplicate chestnut samples (250 g) were collected in
Whirl-Pak bags (NASCO, International lnc.,120 mm x 60 mm) after transport to
the facility (recollection buckets), pre—processing at the facility (receiver in chain
batcher—A), passing through the burner on the elevator batcher-D, passing
through the steamer-H, passing through the counter-rotating rollers and through
the brushes-K, transport on the sorting belt-L, and before packaging at the end of
the peeling line-M (Figure 10 and Figure 11). Samples were transported to the

laboratory on ice and analyzed within 24 h of collection.

Environmental and water samples

Environmental samples were collected during processing (Figure 10 and
Figure 11) on shoot before steamer-E, on shoot after steamer-H, on residue
disposal shoot after brushes-J, from counter-rotating rollers in brushes-l, on shoot
after brushes-K, on sorting belt-L and on shoot after sorting belt-M from 100 cm2
area using sterile kim-wipes (Kimberly-Clark Professional, Rosewell, GA) in
Whirl-Pak bags containing 10 ml of phosphate buffer solution (PBS) (pH = 7.4)

(Figure 10 and Figure 11). 50 ml water samples were also collected from the tube

106

injecting water into the steamer-G, from tube leaking out of steamer after use-F,
residue disposal shoot after brushes-J, after brushes-K and after the sorting belt-
M in 50-ml sterile centrifuge tubes (Corning®, Life Science Group, Lowell, MA,
USA) (Figure 10 and Figure 11). Samples were transported to the laboratory on

ice and analyzed within 24 h of collection.

 

Preharvest

Manual ‘
harvest 1

Transportation ‘ rash chestnu eeled chestn Swab Water
and storage . sample sample sample sample

 

 

 

 

 

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Pre
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(receiver)

  

 

 

Burner $

 

 

Steamer 4

 

 

lBrushes |=

 

 

A
T

 

 

 

 

-——-—a
‘i

 

 

Packaging wk

 

 

 

Figure 11. Sample collection points for fresh and peeled chestnuts. Swab and water
samples were taken during chestnuts peeling.

107

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108

  

Determination of microbial populations in fresh chestnut, peeled chestnut

and environmental samples

Fresh chestnuts were surface sterilized by dipping each chestnut in 99%
ethanol and ﬂaming after which the shell was aseptically removed. Each sample
was weighed (~ 25 9) into a sterile Whirl-Pak stomacher bag diluted 1:5 in
phosphate buffer solution (PBS) (pH 7.4) and homogenized for 1 min (normal
speed) in a stomacher Model 400 (Seward Lab. System, England). The same
procedure, with exception of shell removal was repeated for peeled chestnuts.
The suspension was serially diluted and 100 pl aliquots were inoculated onto
trypticase soy agar (Becton and Dikinson Sparks, Md, USA) plus 0.6% yeast
extract containing 100 ppm of cycloheximide (Sigma-Aldrich, St. Louis, Mo, USA)
(TSA-YEC) for quantiﬁcation of mesophilic aerobic bacteria (MAB) and onto
potato dextrose agar containing 20 ppm streptomycin and 50 ppm ampicillin
(PDA-SA) for enumeration of yeasts and molds.

Environmental swab samples were homogenized for 1 min (normal speed)
in a stomacher Model 400 (Seward Lab. System, England) and then plated for
MAB, yeasts and molds, along with 100 pl aliquots of the waters samples as just
described. MAB plates were counted after 48 h of incubation at 25° C (1 3),
whereas the mold and yeast CFU, were counted after 72 h of incubation at 25° C

(i 3) (APHA, 2001).

109

Identification of microorganisms

Microorganisms isolated from 25 samples (25 g) of symptomatically
thawed peeled chestnuts, containing sticky, yellow ooze on the surface were
picked and subsequently puriﬁed after isolation. Spoilage symptoms were
described, photographed and classiﬁed.

MAB were identiﬁed sequencing rDNA subunits (White, Bruns, Lee, and
Taylor, 1990). Bacterial genomic DNA was extracted using the protocol proposed
by Jacobs et al. (2008). The region including the two spacers (UFLP and URPL)
was ampliﬁed using the method of LiPuma et al. (1999). A total volume of 25 pl
for each reaction contained 60 ng of DNA, 23 pl of AFLPTM ampliﬁcation core mix
(Applied Biosystems, Foster City, Ca, USA), 2.5 units of AmpliTaq Gold DNA
polymerase (Applied Biosystems) and 10 pM of each primer. The primers UFPL
(5'-AG'I‘I'TGATCCTGGCTCAG-3') and URPL (5'-GGTTACCTTGTTACGACTI’-
3') were used for targeting the 16S rDNA region from the kingdom Procaryotae
(bacteria) (LiPuma et al., 1999).

The extracted DNA region was ampliﬁed in a 2720 thermal cycler (Applied
Biosystems). The program included a cycle at 95 °C for 2 min, 30 cycles of 95 °C
for 30 s, 55 °C for 30 s and 71 °C for 1 min, and a ﬁnal elongation at 71 °C for 10
min. The PCR product was placed in wells on an agarose gel (1 .25 %), and
electrophoresed for 3 hours at 70 volts. After electrophoresis, the ampliﬁed region
was puriﬁed using a QiAquick PCR Puriﬁcation kit (Qiagen, Maryland, USA). The
PCR product was sequenced in both directions using moderate throughput _

sequencing (Research Technology and Support Facility, MSU, East Lansing, MI,

110

USA). Sequences were deposited in the Lasergene software (DNA Star Inc,
Madison, WI., USA) and used as queries for similarity searches in the NCBI
nucleotide database (NCBI, 2008). Species reported had a 98 to 100 % match in
both directions.

Yeasts were inoculated on potato dextrose agar, incubated for 72 h at 25°
C (1 3° C) and then sent to the DCPHA (MSU, East Lansing, Mi., USA) for further
identiﬁcation. Yeasts were identiﬁed using the API 200 yeast identiﬁcation
system together with microscopic morphology determinations (Michigan State
University Diagnostic Center for Population and Animal Health (DCPHA)). This
computer-assisted system for rapid identiﬁcation of yeast provides results
comparable to those obtained of conventional morphological methodologies
(Land et al., 1979).

Determining symptoms associated with microorganisms isolated from

spoiled peeled chestnuts

WM

Based on visual observations, the identiﬁed cultures determined to be
associated with spoilage in peeled chestnuts were used in further inoculation
studies. Bacteria were sub-cultured on TSA-YEC and yeasts were sub-cultured
onto PDA-SA and incubated for 48 h at 25 °C. A single colony of each strain was
transferred to 5 ml of Lauria-Bertani (LB) soft media (Becton and Dikinson,
Sparks, Md, USA), incubated for 24 h at 25 °C. and constantly aerated on a
rotator shaker at 150 rpm (model Classic C10, New Brunswick Scientiﬁc Co. Inc,

Edison, NJ, USA). Broth cultures were then sterilely transferred by pouring the

111

subculture into sterile 100 ml LB media. This process was exponentially
repeated, each 24 hour until desired amount of inocula from each organism was

acquired.

Sterilization and inoculation of peeled chestnuts

Whole chestnuts (cv. ‘Colossal’) were mechanically peeled, vacuum-
sealed in low-density polyethylene bags (Kent Buthcers Supply, Grand Rapids,
MI) stored at — 5 °C for ﬁve weeks until used and then were completely thawed
for four hours at 25 °C before use.

The thawed chestnuts were sterilized by two successive 10 min
immersions in 20% sodium hypochlorite (1 L per 0.5 kg of chestnuts) with
constant agitation on a rotator shaker (190 rpm) (model Classic C10, New
Brunswick Scientiﬁc Co. Inc., Edison, NJ, USA) and then subjected to three 15
min washing with sterile-deionized water (1 L per 0.5 kg of chestnuts) with
agitation (190 rpm) to eliminate residual chlorine. Chestnuts were then placed on
sterile blotting paper and air dried in a bio-safety hood with for 25 min.

To conﬁrm chestnut spoilage associated with isolated microorganisms,
individual organisms and equal organism cocktail, containing between 108 to 109
CFU/ml of each organism were used as inocula in the test. Sterilized chestnuts
were placed in ﬂasks contained mixed inocula (1 L per 1 kg of chestnuts) and
agitated on a rotator shaker (200 rpm) for 15 minutes. Inoculated chestnuts were

placed on sterile blotting paper and air dried in a bio-safety hood for 30 min.

112

Treatments of artificially inoculated geeled chestnutsI storage and
microbial analysis

Whole chestnuts (cv. ‘Colossal’) were mechanically peeled, vacuum-
sealed in low-density polyethylene bags (Kent Buthcers Supply, Grand Rapids,
MI) stored at — 5 °C for ﬁve weeks until used and then were completely thawed
for four hours at 25 °C before use. Eight treatments were implemented in an
attempt to mimic industrial processes (Table 5). All liquid-solution treatments,
except heat treatment (hot water), X-ray irradiation and ozone solutions, were
thoroughly mixed, to achieve the desired concentration in 30 L of water.
Inoculated ~500 g of chestnuts were immersed in each solution for 2 min. After
treatment, three ~1509 (~12 chestnuts) samples per treatment were collected.

For hot water treatments, peeled chestnuts were treated with heated water
generated by a laboratory water bath (model DurabathT", Baxter Scientiﬁc,
Melrose Park, IL). After the desired temperature was reached, inoculated
chestnuts were immediately submerged in the water for 2 min. Temperature was
constantly monitored using a 7 mm immersion thermometer (Fisherbrand
Scientiﬁc by Ertco, UK). Five hundred grams of inoculated chestnuts were dipped
at once in warm water for 2 min. After treatment, three ~150 9 (~12 chestnuts)
samples per treatment were collected

Ozone was generated using a commercial ozone generator (Aqua Air

Technologies, Inc., Bloomﬁeld, MI) equipped with an oxygen concentrator model
CD 1OIAD Corona Discharge ozone generator system (Clear Water Tech., Inc,

San Luis Obispo, CA). Gas was delivered through an inlet line directly into the

113

water, which was constantly circulated through a 30 L water container. Water
containing up to 0.7 ppm ozone was obtained within 25 min. After the desired
concentration was achieved, ~500 g of inoculated chestnuts were immersed in
the ozone solution for 2 min. The ozone concentration was constantly monitored
using a portable dissolved ozone meter (model OZ-21P, DKK-TOA Corporation,
Takadanobaba, Shinjuku-ku, Tokyo, Japan).

Inoculated peeled chestnuts were irradiated at target doses of 0.5, 1.0,
1.5, and 2.0 RC using a low-energy X-ray food irradiator (Rayfresh Foods Inc,
Ann Arbor, MI). The sample consists of a single layer of chestnuts (~12
chestnuts, ~150 g) in a Whirl-Pak plastic bag. Incident dose was measured at six
locations on the surface using radiochromic ﬁlm dosimeters (GAF3001DS, GEX
Corp., CO). To maximize dose uniformity on the surface a dual tube and double
treatment technique was used. These means that target doses of low-energy X-
ray, were unifonnly distributed by two-tubes to the ﬂat surface of each side of the
chestnut sample by manually turning the sample once.

All populations of MAB and Y were determined on the day of treatment

(day 0), and after 10 and 18 days of storage at 4° C.

114

Table 5. Post-processing treatments used In artificially inoculated peeled chestnuts

 

 

 

 

 

 

 

 

 

 

 

Commercial name Active imedient Concentration
X-ray irradiation (Rayfresh Foods low-energy X-ray irradiation 0.5, 1, 1.5, 2 kGy
lnc., Ann Arbor, MI, USA)
StoroxTM (BioSafe Systems, 27% Hydrogen dioxide + 2% 2,700-ppm
Glastonbury, CT, USA) peracetic acid hydrogen dioxide

+ 200-ppm
peracetic acid
Agri-Cide (Life Science Group, 18.25 — 21.75% Copper sulfate 1 ppm
Inc. Monticello, IN, USA) pentahydrate
Chlorine dioxide solution (ICA c|02 + H20 10 ppm
TriNova, LLC Forest Park, GA)
Ozone solution (Aqua Air 03 4» H20 0.70 ppm
Technologies, Inc., Bloomﬁeld,
NL USA)
Peracetic Acid (Lenntech, Peracetic acid 80 ppm
Rotterdamseweg, Netherlands)
Chlorine solution (Champion 6% Sodium hypochlorite 100 ppm
packaging, Inc. Woodridge, IL,
USA)
Sodium chloride (Sigma-Aldrich, Sodium chloride 0.2 M
St. Louis, MO, USA)
Hot water Heat 65 °C
Statistical analysis

WWW
One-factor analysis of variance (ANOVA) was done on all microbial
populations obtained from peeled chestnuts. Multiple analysis of variance
(MANOVA) was used to determine differences among the microbial populations
from different sampling points. Signiﬁcance among the means was determined by
the Tukey post-hoc multiple comparisons of means test at the 95% family-wise
conﬁdence level (p=0.05). Calculations were performed by using the “R: A
language and environment for statistical computing” statistical package (R

Development Core Team, 2007) (Ott and Longnecker, 2001).

115

WWW

Repeated measurement design with analysis of variance (ANOVA) was
done to average microbial count data (MAB and yeast) obtained from treated
peeled chestnuts. Since the assignment of samples to the different treatment
conditions as well as the choice of subsamples at each point of measurement
were completely at random, sphericity can be assumed making multivariate as
well as degrees of freedom corrections unnecessary (Keselman, Algina, and
Kowalchuk, 2001; Ott and Longnecker, 2001). This procedure will indicate if a
treatment signiﬁcantly reduce the microbial population during 18 days of storage.
Signiﬁcance among the means was determined using the Tukey post-hoc
multiple comparisons of means test at the 95% family-wise conﬁdence level
(p=0.05. These calculations were performed using the same statistical package

listed above.

Results

Microorganisms associated with spoiled peeled chestnuts

Two bacteria (Rahnella sp. and Curtobacterium sp.), and one yeast
(Candida sp.) were consistently isolated from the 25 samples (~ 25 g) of vacuum-
packaged, peeled chestnuts thawed and stored for 10 to 15 days at 4 °C. These
three microbes were considered the primary cause of spoilage as they each
produced yellow, sticky ooze when artiﬁcially inoculated on sterile chestnuts
(Figure 12-a to 12-d). Rahnella sp. and Candida sp. were also found on fresh

chestnuts in an earlier study (Chapter 2 of this thesis) and therefore the source of

116

contamination of peeled chestnuts may be fresh chestnuts that are naturally
contaminated when brought from the field. Although some browning was also
associated with the spoilage, this can occur without the development of ooze as
the chestnuts oxidize in time.

In most cases, all three spoilage organisms were present in spoiled
chestnut; however, each alone, was also capable of causing spoilage. No
obvious visual difference in terms of symptoms produced if either one or more
organisms causing spoilage were observed (Figure 12-b to 12-d). The chestnuts
were visually spoiled when the MAB and yeast populations exceeded 8 log

CFU/g.

 

 

 

 

 

 

 

Figure 13. Peeled chestnuts, 10 days after inoculation and stored at 4 °C. a) Non-
Inoculated sterilized control, b) inoculated with Curtobacterium sp., c) inoculated with
Rahnella sp., d) inoculated with Candida sp. Image in this thesis is presented in color.

117

Microbial population

Chestnut samples

Chestnut samples were collected at different locations before, during and
after peeling, to determine how spoilage organisms were becoming established
on the processed peeled product. Populations of MAB, molds and yeast before
processing (buckets and receiver) varied from 2.50 to 2.90 logs CF U/g. After
peeling, MAB (F (6,2) = 362.19, p < 0.01) and yeast (F (6,2) = 8.66, p < 0.01)
increased signiﬁcantly up to 5.39 and 3.09 logs CFU/g, respectively. In contrast,
mold populations at the end of processing decreased to <1.7 logs CF Ulg (Figure
13). A signiﬁcant difference was seen between all microbial populations within
samples collected before, during and after processing (approx F (6,2) = 19.59, p
< 0.01). In general, the brushes played an important role in signiﬁcantly

increasing MAB and yeast populations in peeled chestnuts.

118

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

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Sampling point

Figure 14. Total MAB, molds and yeast counts (log cfu g'1) on peeled chestnuts before and

during peeling process (Brulage peeler - Boema; Neive, Italy). 1Values followed by the same
letter within organisms are not signiﬁcantly different at p = 0.05 (ANOVA) (Tukey multiple
comparison of means). Minimum detectable level (MDL) = Minimum possible count (0.5) x
minimum dilution factor x inoculated aliquot (100 pl). Error bars indicate standard deviation.

W

Environmental samples were taken from points along the chestnut peeling
line to help determine the point of chestnut contamination. Based on
environmental swab samples, populations of MAB and yeast differed signiﬁcantly
along the peeling line. MAB, molds and yeasts were undetectable in the steamer
and before brushes (< 2.7 logs CFU/cm"). However, after the sorting belt and
before packaging MAB (F (5,2) = 243.06, p < 0.01) and yeast (F (5,2) = 15.05, p

< 0.01) populations increased signiﬁcantly to 5.60 and 4.72 logs CPU/100 cmz,

119

respectively. In contrast, mold populations after processing were <2.7 logs
CFU/100 cm2 (Figure 14). A signiﬁcant difference was seen between all microbial
populations within environmental samples collected during the peeling process
(approx F (5,2 = 13.41, p < 0.01). These reafﬁrms that the brushes played an

important role in signiﬁcantly increasing MAB and yeast populations.

6

4

Log (CFUI10 cm?)

 

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Steamer Before Brushes Brushes After Brushes Sorting Belt After Sorting Belt
Sampling point

 

Figure 15. Total MAB, molds and yeast counts (log cfu cm'z) on environmental samples
during peeling process (Brulage peeler - Boema; Neive, Italy). 1Values followed by the same
letter within organisms are not signiﬁcantly different at p = 0.05 (ANOVA) (Tukey multiple
comparison of means). Minimum detectable level (MDL) = Minimum possible count (0.5) x
minimum dilution factor x inoculated aliquot (100 pl). Error bars indicate standard deviation.

120

Misti—males

Water samples were taken from points along the chestnut peeling line to
help determine the point of contamination. MAB, molds and yeasts were
undetectable in the steamer and after the steamer (< 1.7 logs CFU/ml).
However, after the brushes, MAB (F (4,2) = 1136.36, p < 0.01) and yeast (F (4,2)
= 10.59, p < 0.01) signiﬁcantly increased to 3.73 and 4.98 logs CFU/ml,
respectively. The mold population remained constant during the entire process
(<1.7 logs CFU/ml) (Figure 15), but above population levels found in the steamer.
A signiﬁcant difference was seen between microbial populations within
environmental samples collected during the peeling process (approx F (4,1) = '
428.37, p < 0.01). In general, a signiﬁcant increase in MAB and yeast was again

observed in water collected from brushes.

121

 

 

 

 

   

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

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Steamer After Steamer Brushes After Brushes

Sampling point
Figure 16. Total MAB, molds and yeast counts (log cfu g'1) in water samples during peeling
process (Brulage peeler - Boema; Neive, Italy). 1Values followed by the same letter within
organisms are not signiﬁcantly different at p = 0.05 (ANOVA) (Tukey multiple comparison of

means). Minimum detectable level = Minimum possible count (0.5) x minimum dilution factor x
inoculated aliquot (100 pl). Error bars indicate standard deviation.

Post-processing treatments

Sanitizers and hot water

To determine if the shelf life of chestnuts could be extended after thawing,
several microbial reduction strategies were assessed using chestnuts inoculated
with the three identiﬁed spoilage organisms. The mean plus or minus the
standard deviation for populations of MAB and yeast were evaluated immediately

after each treatment, and after 10 and 18 days of storage at 4° C (Figure 16).

122

85

.......................................

/" ' w Control (Water)- c1
’ / 4.x - Ozone solution-c
,' V :1» NaCl-c
I ' .-. Sodium hypochlorite- bc
- Chlorine dioxide soln.- b

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to? _ A 18

 

Storage (Days)
Figure 17. Mean MAB and yeast counts (log cfu g") from peeled chestnuts treated with
different sanitizers during 18 days of storage at 4' C. 1Overall data point followed by the same
letter within treatments are not signiﬁcantly different at p = 0.05 (Repeated measurement design —
ANOVA with post-hoc Tukey multiple comparison of means). Error bars indicate standard
deviation.

Two minutes exposure time to 10 ppm Chlorine dioxide, 1 ppm copper
sulfate pentahydrate and 80 ppm peracetic acid signiﬁcantly reduced MAB and
yeast population during 18 days of storage in comparison with the non-treated
control (F (8,2) = 43.89, p < 0.01). However, on day 18 the microbial population
had already surpassed the desired levels and spoilage could be observed. Only 2
min exposure time to 2,700-ppm hydrogen dioxide + 200-ppm peracetic acid and
hot water treatment signiﬁcantly reduced the mean MAB and yeast population
during 18 days of storage in comparison with the non-treated control (F (8,2) =

43.89, p < 0.01). In both cases, the mean microbial population on day 18 was

123

signiﬁcantly lower by 1.24 and 1.55 logs CF U/g respectively, in comparison with

the non—treated control and no spoilage could be observed.

X-raz irradiation

To determine if spoilage could be managed and shelf life extended after
thawing, various X-ray irradiation doses were tested on chestnuts inoculated with
the three microbes associated with spoilage. The mean of MAB and yeast
populations from the various doses of X-ray irradiation treatment were
determined immediately after treatment, and 10 and 18 days of storage at 4° C

(Figure 17).

124

./ ° -- Control -e1
_:/: 0.5 kGy- d
-- 1.0kGy-c

1 --1.5kGy-b
- 2.0kGy-a3

 

 

o 10 '7 ‘ 13
Storage (Days)
Figure 18. Mean MAB and yeast counts (log cfu g") from peeled chestnuts treated with
different X-ray Irradiation doses during 18 days of storage at 4° C. Overall data point
followed by the same letter within dosis are not signiﬁcantly different at p = 0.05 (Repeated

measurement design — ANOVA with post-hoc Tukey multiple comparison of means). Error bars
indicate standard deviation.

All X-ray irradiation doses signiﬁcantly reduced the MAB and yeast
populations during 18 days of storage in comparison with the non-treated control
(F (4,2) = 138.97, p < 0.01). The mean microbial reductions when compared with
the non-treated control on day 18 for 0.5, 1.0, 1.5 and 2.0 kGy were 1.47, 2.33,
3.69 and 4.02 logs CFU/g, respectively. After 18 days no spoilage could be

observed in any of the treated chestnuts.

Discussion

Within 12 days after thawing, peeled chestnuts developed a sticky, yellow

ooze over the surface that affected the ﬁnal quality, acceptability and shelf life.

125

The three primary spoilage microorganisms identiﬁed included two bacteria
Rahnella sp., Curtobacterium sp., and one yeast, Candida sp. There is no
evidence indicating that ﬁlamentous fungi play an important role during spoilage.
Contamination of peeled chestnuts was strongly inﬂuenced by the peeling
process. The traditional method of peeling chestnuts, (e.g. manual elimination of
the shell and pellicle from fresh chestnuts) is laborious and requires heat and
knives. In Michigan, peeled chestnuts are obtained by processing fresh chestnuts
with a commercial-level peeling system (Boema; Neive, Italy) (Appendix B) after
which the chestnuts are usually vacuum-packed and frozen (Guyer, et al., 2003).
Other mechanical methods, such as air-impingement de-sheiling, are also
available but are not used in Michigan (Gao, et al., 2008). Michigan chestnut
samples, and environmental and water microbial surveys indicated that average
MAB and yeast populations signiﬁcantly increased during peeling with the skin
- separator (brushes) and sorting belt identiﬁed as key points for contamination.
Similar results have been reported for other commodities and processing
plants. Monitoring of several organisms, like Listeria monocytogenes during
produce processing has indicated that cross-contamination from contaminated
zones in the processing line, ﬂoor surface and gloves is critical and extremely
difﬁcult to control (Pappelbaum, et al., 2008). Others have reported that utensils,
processing tables, and lines may also increase the levels of MAB, colifonns and
pathogenic organisms like Escherichia coli O157:H7 (Christison, Lidsay, and Von
Holy, 2008). All of these studies have strongly indicated that the processing
environment may play an important role in maintaining and enhancing pathogen

populations. Because of these possibilities, and since MAB and yeast populations

126

signiﬁcantly increased by more than 2 logs in chestnuts between harvest and
after peeling, improved and more effective microbial reduction strategies after
and during processing are needed to ensure the quality and prolong the shelf life
of peeled chestnuts.

An alternative method to reducing microorganisms in the ﬁnal product is to
eliminate microbial presence by constantly disinfecting sections from the
processing line, using different detergents, sterilizing agents and enzymes. These
methods have been evaluated in dairy and meat processing lines (Manvi and
Anand, 2001; Anon., 1970) and have also proven to be effective in removing
microbial bioﬁlms from dispensing equipment (Walker, Fourgialakis, Cerezo, and
Livens, 2007). Neverhteless, further studies must be done to apply these
methods to the chestnut peeler

Another alternative is to try to inactivate these spoilage microorganisms
after processing and prior to storage. In most cases, sanitizing agents are added
to processing water and consequently to the product, to reduce microbial
populations and prevent cross-contamination of the processed commodity
(Sapers, 2003). Hydrogen dioxide concentration of 1 to 10 percent have been
used to reduce the microbial population and extend the shelf life of whole fruits,
like blueberries, fresh-cut fruits and vegetables (Crowe, Bushway, and Bushway,
2005; Sapers, 2003). Hydrogen dioxide is Generally Recognized As Safe
(GRAS), because it is reduced to water and oxygen after treatment (Sapers,
2003). Although treatments containing high concentrations of hydrogen dioxide
may be useful in reducing microbial populations and extending shelf life, the use

of these treatments in commodities like lettuce, blueberries and others, might be

127

limited because of possible plant surface oxidation resulting in product damage
(Aharoni, Copel, and Fallick, 1994; Block, 1991; Simmons, 1997; Ukuku and
Sapers, 2001; Ukuku et al., 2001). The present study indicated that hydrogen
dioxide (1:100 dilution of StoroxTM, BioSafe Systems, Glastonbury, CT)
signiﬁcantly reduced the mean populations of MAB and yeast in peeled
chestnuts, during 18 days of storage in comparison with the non-treated control.

Warm water (65 °C) was found to be among the best methods in
signiﬁcantly reducing spoilage as well as MAB and yeasts in peeled chestnuts,
during 18 days of storage in comparison with the non-treated control. Heat
treatments are often applied for a relatively short time (Fallik, 2004), as hot water
clips or rinsing, vapor heat, hot dry air (Fallik, 2004) and other relatively new
developed technologies like far infrared radiation (Tanaka et al. 2007).
Historically, water has been the preferred medium for most applications since it is
a more efﬁcient heat transfer medium than air (Fallik 2004). Immersion of
avocado fruit (cv. ‘Hass’) for 20—30 min at 40-42 °C controls decay and
enhances the storage quality (Fallik, 2004). Immersion of sweet chestnuts in 45
°C water for 45 min enhanced storage quality of fresh product (Jerimini, et al.,
2006). A ﬁve log reduction of Escherichia coli O157:H7 was also observed after
immersing apples in 80 and 90 °C water for 15 seconds (Fallik, 2004).

Studies have demonstrated that irradiation may also be capable of
reducing pathogens and other microorganisms more than ﬁve logs in many
foods. irradiation is approved by the Food and Agriculture Organization
(FAO)/lnternational Atomic Energy Agency (lAEA)NVorld Health Organization

(WHO), but irradiated products must be labeled properly using the Radura

128

symbol. Given the lack of evidence for toxic residues irradiation may become an
attractive alternative to chemical sanitizers. However, irradiation remains
expensive, requires special facilities and still lacks consumer acceptability (Diehl,
1995; Gamage, Faith, Luchansky, Buege, and lngham, 1997; Howard, Miller, and
Wagner, 1995; Kilcast, 1995; Luh, 1997; Niemira, 2003; Saroj, et al., 2006; Smith
and Pillai, 2004). All evaluated X-Ray irradiation doses (0.5, 1, 1.5 and 2 kGy)
signiﬁcantly reduced the mean population of MAB and yeast during 18 days of
storage and decreased spoilage in comparison with the non-treated control.
Using the highest dose of radiation, population of MAB and yeast were 4 log
CFU/g lower after 18 days of storage. Similar results have been reported for
other commodities (Niemira, 2003). Nevertheless, it is important to consider that,
both hot water treatments and irradiation may damage produce tissue and leak
nutrients by breaking down cell wall material, producing off-ﬂavors in the treated
produce (Lydakis and Aked 2003; Niemira, 2003; Saroj, et al., 2006; Soto-
Zamora, 2005). There was no evidence indicating that the peeled chestnuts were
negatively affected by any of the previously mentioned treatments, but further
sensory evaluation is recommended to discard possible effects.

In conclusion, the present study showed that two bacteria Rahnella sp.,
and Curtobacterium sp. and the yeast Candida sp. were the primary causes of
spoilage of mechanically peeled chestnuts. Overall, MAB and yeast in chestnuts,
water and environmental samples signiﬁcantly increased during peeling, with the
skin separator (brushes) and sorting belt being key points for contamination.
Nevertheless, several post-processing treatments were able to inhibit the growth

of these microorganisms. Among these, 2,700-ppm hydrogen dioxide + 200-ppm

129

peracetic acid, immersion in hot water and X-ray irradiation were most effective.
The prospect of adopting these methods as commercial post-processing
treatments needs to be examined and may have several advantages for the
peeled chestnut industry. Cost, efﬁcient application, monitoring and
implementation of each method must also be examined to maximize economic
gains. All of the evaluated treatments are presently used in food processing and
are generally regarded as safe at the appropriate concentration. These may be
implemented as part of a sustainable integrated pest management strategy for

peeled chestnut spoilage.

130

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137

APPENDICES

138

APPENDIX A

MYCOTOXINS IN FRESH MICHIGAN CHESTNUTS

139

Figure 19. Presence of mycotoxins In two cultivars of fresh Michigan chestnuts. Five, 350 g
of healthy kemei samples per cultivar were processed to extract mycotoxins and quantiﬁed by
competitive direct ELISA test (Neogen Corporation, Acumedia Lansing, MI, USA) as speciﬁed by
the manufacturer. 1Data points followed by the same lower caseletter within the same day are not
signiﬁcantly different at p = 0.05 (T-test). 2Overall data point followed by the same capitalized
letter within chestnut part-harvesting method are not signiﬁcantly different at p = 0.05 (Repeated
measurement ANOVA). Error bars indicate standard deviation.

140

Concentration (ppb) Concentration (ppm)

Concentration (ppb)

0

100 200 300 400 500

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Storage (Days)

141

APPENDIX B

COMMERCIAL-LEVEL PEELING SYSTEM (BOEMA; NEIVE, ITALY)

142

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143

APPENDIX C

CHESTNUT FRUIT MORPHOLOGY

144

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145

APPENDIX D

WORLD CHESTNUT PRODUCTION

146

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147

APPENDIX E

INHIBITION PROPERTIES OF PELLICLE, HILUM, SHELL AND KERNEL
AGAINST Cladosporium cucumerinum

148

 

Figure 23. Inhibition properties of pellicle, hilum and shell against Cladosporium
cucumerinum. To determine the inhibition properties of the different parts of the chestnuts (shell,
hilum, pellicle and kernel), 70 percent acetone extract from these parts from two different cultivars
were evaluated (‘Everfresh' = C. mollissima and ‘Colossal’ = C. sativa x C. crenata). Paper disc
agar diffusion method was employed, using ﬁlter paper discs of 10 mm diameter (Whatman,
Shleicher and Schuell International Ltd.) impregnated with different concentration of the extracts.
After the discs were put onto the agar, the organism was surface inoculated by spraying 10 ml of
a sterile distilled-water solution containing between 220-300 spores/ml of the pathogen. The
plates were sealed by the use of paraﬁlm and stored inverted at room temperature (25° C 1 3°).
A: 50 ul-sheIl-‘Colossal', B: 100 uI-shell-‘Colossal’, C: 100 uI-pellicle-‘Colossal’, D: 50 pI-pellicle-
‘Colossal’, E: 100 pl-hilum-‘Colossal', F: 10 uI-pellicIe-‘Everfresh', G: 5 ul-hilum-‘Everfresh', H: 100
pl-kerneI-‘Everfresh’, I: 100 uI-pellicle-‘Everfresh‘, J: Control (70 percent acetone). K: 5 uI-hilum-
‘Colossal', K: 100 pl-pellicle-‘Everfresh', M: 50 pi-pellicle-‘Everfresh’, N: 50 pl-pellicIe-‘Colossal',
0: Control (70 percent acetone), P: 50 pl-shell-‘Colossal', Q: 100 pl-kernel-‘Colossal’, R: 10 pl-
pellicie—‘Everfresh‘. Different levels of inhibition can be observed depending on the cultivar and
different parts of the chestnuts. ‘Everfresh' had higher levels of inhibition in comparison with
‘Colossal’. The pellicle presented the highest zone of inhibition, followed by the shell and the
hilum. The kernel did not inhibit the growth of the pathogen. Image in this thesis is presented in
color.

149

   

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