The Pennsylvania State University

The Graduate School

Department of Agricultural and Biological Engineering

ALTERNATIVE MUSHROOM PRODUCTION SYSTEMS USING NON-

COMPOSTED GRAIN-BASED SUBSTRATES

A Thesis in

Agricultural and Biological Engineering

by

Mark A. Bechara

©2007 Mark A. Bechara

Submitted in Partial Fulfillment

of the Requirements

for the Degree of

Doctor of Philosophy

December 2007

The thesis of Mark A. Bechara was reviewed and approved* by the following:

Paul H. Heinemann Professor of Agricultural and Biological Engineering Thesis Advisor Chair of Committee

Paul N. Walker Professor of Agricultural and Biological Engineering

Tom L Richard Associate Professor of Agricultural and Biological Engineering

John M Regan Assistant Professor of Environmental Engineering

C. Peter Romaine Professor of Plant Pathology

Roy E. Young Professor of Agricultural and Biological Engineering Head of the department of Agricultural and Biological Engineering

*Signatures are on file at the Graduate School

ii ABSTRACT

The commercial production system of Agaricus bisporus mushroom relies entirely on composting as a means to generate a mushroom-specific substrate, and all aspects of this traditional system are designed for the preparation, processing, and handling of this compost-based substrate. Furthermore, the substrate, which is a mixture of compost and delayed-release supplements, has been refined for maximum mushroom production, and although widely adopted by mushroom producers around the world, this system is environmentally problematic (odor emissions, nutrient-rich run-off, and substrate disposal limitations) and these problems will only intensify with urban sprawl.

In this study, grain-based mushroom production systems were evaluated as a potential alternative to the traditional compost-based system for A. bisporus. Two types of grain-based systems were developed and called– “Satellite Mushroom Production

System” (SMPS), and “Complete On-site Mushroom Production System” (COMPS).

Mushroom producers adopting the SMPS, would use a substrate primarily composed of commercial grain spawn, whereas mushrooms producers adopting the COMPS would use a subsrate composed of cereal grains mixed with oilseed (grain/oilseed). Both systems were tested and refined for mushroom production and substrate bioefficiency (BE).

Furthermore, the theoretical designs and the direct costs for mushroom production for each system were developed and calculated.

For the SMPS, the factors influencing mushroom production were: adding a layer of water-holding materials below the commercial grain spawn substrate, heat-sterilizing casing with the addition of activated carbon, and type and level of delayed-release supplement used. Overall, the highest yield of mushrooms for a commercial grain spawn

iii substrate was 14.28 kg/m2 with a corresponding BE of 177% and was obtained for a

treatment containing 5% S41- an underlying layer of perlite (2000 ml) and cased with a

heat-sterilized casing containing 25% activated carbon. The design of SMPS entails

different steps that start with mixing of the substrate ingredients, transferring the

substrate to trays, and subsequent steps are comparable to traditional compost-based

systems. Based on the cost model developed for the SMPS, the cost of production/unit

weigh of mushroom is $4.00/kg. This is much higher than the market cost of A. bisporus

and mushroom producers would not make a profit.

For the COMPS, the factors that were tested for mushroom production using

grain/oilseed substrate were type of cereal grain used, type and rate of oilseed added, pre-

conditioning the substrate with Scytalidium thermophilum (a mushroom compost

thermophile), and moisture content in the grain portion of the substrate. Furthermore, two

additional mushroom producing species (A. blazei and Pleurotus eryngii) were grown on the grain/oilseed substrate to determine whether the grain-based system could be extended to the production of other mushroom forming species.

For A. bisporus, adding oilseeds to a basal substrate composed of millet grain increased yield and a 15% amendment of soybean gave the highest yield (16.9 kg/m2 with a corresponding BE of 205%). Furthermore, preconditioning the substrate with S. thermophilum improved yield on oat based-substrates and decreased spawn-run time for all grain/oilseed substrates. Moisture contents ranging from 50-65% seem adequate for mushroom production, whereas the peak in yield is observed between the 55% and 60% moisture levels.

iv For A. blazei and P. eryngii mushroom production, grain/oilseed substrate

successfully supported mushroom production. For A. blazei, millet with 30% niger

yielded 15.9 kg/m2, whereas, an oat/oilseed substrate provided the highest yield of 106 g

for P. eryngii. Overall, the highest yield for A. bisporus was observed for a millet/ 5% soybean substrate with an additional amendment of 5% delayed-release supplement

(Promycel Target®) which produced 21.3 kg/m2 with a BE of 273%. The system design

is composed of an aseptic processing unit that converts the raw materials into a suitable

mushroom substrate, and steps following the processing of the grain/oilseed substrate are

comparable to the SMPS. Based on the cost model developed for the COMPS, the cost of

production/unit weight of mushrooms is $1.40/kg, and this would provide mushroom producers with a profit.

Based on findings from this research, developing an alternative commercial A.

bisporus mushroom production system using grain-based substrates is a highly promising

alternative to commercial compost-based system and its environmental problems. Yield from grain-based substrates is still lower than compost-based substrates but further refinement of the substrate ultimately will increase mushroom yield. A pilot-scale system is needed to test scale-up of this system for mushroom production on grain-based substrates.

v TABLE OF CONTENTS

LIST OF FIGURES...... X LIST OF TABLES...... XII ACKNOWLEDGMENTS...... XIV 1. INTRODUCTION ...... 1 2. LITERATURE REVIEW ...... 5 2.1. AGARICUS BISPORUS MUSHROOM FUNGUS ...... 5 2.2. SPAWN PRODUCTION ...... 6 2.3. NUTRIENT SUPPLEMENTS USED BY MUSHROOM PRODUCERS ...... 8 2.4. COMMERCIAL PRODUCTION OF AGARICUS BISPORUS ...... 9 2.4.1. Composting in Commercial Agaricus bisporus Production...... 9 2.4.2. Mushroom Fungus Spawning...... 12 2.4.3 Substrate Casing and Casing Inoculum...... 13 2.4.4. Mushroom Fungus Pinning and Harvesting ...... 15 2.4.5. Metabolism and Physiology of A. bisporus...... 15 2.5. AGARICUS MUSHROOM PRODUCTION ON NON-COMPOSTED SUBSTRATES ...... 19 2.6. AGARICUS BISPORUS USED IN ANIMAL FEED ...... 21 2.7. SOLID STATE FERMENTATION ...... 22 2.7.1. Definition of Solid State Fermentation...... 22 2.7.2. Support Material Used in SSF ...... 23 2.7.3. Grains and Oilseeds...... 24 2.7.4. Environmental and Process Parameters in SSF ...... 26 2.7.5. Fungal Biomass Estimation in SSF...... 28 2.8. ASEPTIC PROCESSING OF PARTICULATE MATERIALS...... 30 2.8.1. Gelatinization of Starch ...... 30 2.8.2. Thermal Processing and its Application to Food...... 31 2.8.3. Steam-based Thermal Sterilization Processes ...... 31 2.8.4. Aseptic Processing History and Description ...... 32 2.8.5. Elements of a Continuous Aseptic Processing System ...... 34 2.8.5.1. Heating Section...... 34 2.8.5.2. Holding Section...... 34 2.8.5.3. Cooling Section...... 34 2.8.5.4. Aseptic Filling and Packaging Section ...... 35 2.8.6. Liquid-based Segmented Flow Aseptic Processing System...... 35 2.8.7. Steam-based Segmented-flow Processing System ...... 36 2.9. CONTINUOUS PRODUCTION OF MUSHROOM SPAWN ...... 37 2.10. SUMMARY OF LITERATURE REVIEW ...... 38 CHAPTER 3 PRODUCTION OF AGARICUS BISPORUS MUSHROOMS ON COMMERCIAL GRAIN SPAWN MIXED WITH S41 AND S44 SUPPLEMENTS...... 39 3.1 ABSTRACT ...... 39 3.2. INTRODUCTION ...... 40 3.3. METHODS ...... 44 3.3.1. Mushroom Grain Spawn, Nutrient Supplement and Water-holding Material ...... 44 3.3.2. Preparation of Mushroom Production Containers ...... 44 3.3.3. Mushroom Production Chamber...... 45 3.3.4. Summary of the Experimental set-up ...... 45 3.3.5. Parameters Evaluated and Analysis of Data ...... 46 3.4. RESULTS ...... 46 3.5. DISCUSSION ...... 49 3.6. CONCLUSIONS...... 52

vi CHAPTER 4 FACTORS INFLUENCING MUSHROOM YIELD IN NON-COMPOSTED COMMERCIAL GRAIN SPAWN SUBSTRATES...... 54 4.1. ABSTRACT ...... 54 4.2. INTRODUCTION ...... 55 4.3. METHODS ...... 58 4.3.1. Mushroom Hybrid Nutrient Supplement and Water-holding Material ...... 58 4.3.2. Preparation of Mushroom Production Containers ...... 58 4.3.3. Environmental Conditions in Mushroom Production Rooms ...... 59 4.3.4. Experimental Set-up and Analysis of Data ...... 60 4.4. RESULTS ...... 61 4.5. DISCUSSION ...... 67 4.6. CONCLUSIONS...... 68 CHAPTER 5 EFFECT OF DELAYED-RELEASE SUPPLEMENTS IN GRAIN-BASED SUBSTRATE ON YIELD OF THE MUSHROOM (AGARICUS BISPORUS) ...... 70 5.1. ABSTRACT ...... 70 5.2. INTRODUCTION ...... 71 5.3. METHODS ...... 74 5.3.1. Mushroom Grain Spawn Supplements and Water-holding Material ...... 74 5.3.2. Preparation of Mushroom Production Containers ...... 74 5.3.3. Settings in Mushroom Tray Reactor ...... 75 5.3.4. Summary of the Experimental Design...... 75 5.3.5. Parameters Evaluated and Analysis of Data ...... 75 5.4. RESULTS ...... 76 5.4.1 Mushroom Yield...... 76 5.4.2. Substrate Bioefficiency...... 79 5.4.3. Mean Mushroom Weight...... 79 5.4.4. Substrate Temperature...... 79 5.5. DISCUSSION ...... 81 5.6. CONCLUSIONS...... 83 CHAPTER 6 EVALUATING THE ADDITION OF ACTIVATED CARBON TO HEAT TREATED MUSHROOM CASING FOR GRAIN-BASED AND COMPOST-BASED SUBSTRATES...... 85 6.1. ABSTRACT ...... 85 6.2. INTRODUCTION ...... 86 6.3. METHODS ...... 89 6.3.1. Spawn, Nutrient Supplement and Substrate ...... 89 6.3.2. Casing Preparation...... 89 6.3.3. Preparation of Mushroom Trays...... 90 6.3.4. Conditions in Tray Bioreactor ...... 90 6.3.5. CO2 Absorption Test ...... 91 6.3.6. Experimental Design...... 92 6.4. RESULTS ...... 93 6.5. DISCUSSION ...... 98 6.6. CONCLUSIONS...... 102 CHAPTER 7 CULTIVATION OF AGARICUS BISPORUS AND AGARICUS BLAZEI ON SUBSTRATES COMPOSED OF CEREAL GRAINS AND OILSEEDS...... 104 7.1 ABSTRACT ...... 104 7.2. INTRODUCTION ...... 105 7.3. MATERIALS AND METHODS...... 109 7.3.1. Fungal Strains, Seeds, and Supplements...... 109 7.3.2. Grain Substrate Preparation Process...... 109 7.3.3. Mushroom Production Tray Preparation ...... 110 7.3.4. Tray Bioreactor Conditions ...... 111

vii 7.3.5. Measured Variables in Tray Bioreactors...... 111 7.3.6. Experimental Set-up and Statistical Analysis...... 112 7.4. RESULTS ...... 113 7.4.1. Production Time...... 113 7.4.2. Environmental Conditions in Tray Bioreactors ...... 113 7.4.3. Agaricus bisporus ...... 115 7.4.4. Agaricus blazei...... 119 7.5. DISCUSSION ...... 121 7.6. CONCLUSIONS...... 124 CHAPTER 8 PRE-INCUBATING NON-COMPOSTED GRAIN SUBSTRATES WITH THE THERMOPHILIC FUNGUS SCYTALIDIUM THERMOPHILUM FOR MUSHROOM (AGARICUS BISPORUS) PRODUCTION...... 126 8.1. ABSTRACT ...... 126 8.2. INTRODUCTION ...... 127 8.3. METHODS AND MATERIALS ...... 130 8.3.1. Fungal Cultures Substrate Materials...... 130 8.3.2. Preparation of Grain Substrates...... 131 8.3.3. Preparation of Mushroom Production Containers ...... 131 8.3.4. Environmental Conditions in Mushroom Production Chamber ...... 132 8.3.5. Experimental Set-up and Analysis of Data ...... 133 8.3.6. Oxygen Uptake Rate Evaluation ...... 135 8.4. RESULTS ...... 136 8.4.1. Preliminary Incubation experiments with S. thermophilum...... 136 8.4.2. Treatments with Shorter Incubation Durations with S. thermophilum ...... 137 8.4.3. Oxygen Uptake Rate Evaluation ...... 139 8.5. DISCUSSION ...... 140 8.6. CONCLUSIONS...... 142 CHAPTER 9 EFFECT OF SUBSTRATE MOISTURE CONTENT ON AGARICUS BISPORUS OXYGEN CONSUMPTION RATE, HYPHAL EXTENSION RATE, MUSHROOM YIELD, AND BIOEFFICIENCY...... 144 9.1. ABSTRACT ...... 144 9.2. INTRODUCTION ...... 145 9.3. METHODS ...... 148 9.3.1. Fungal Cultures and Substrate Materials...... 148 9.3.2. Substrate Preparation for OUR Measurements and Mushroom Fruiting...... 149 9.3.3. Procedure for OUR Measurement ...... 150 9.3.4. Oxygen Uptake Rate (OUR) Calculation...... 151 9.3.5. Dry Matter and Ash Measurements ...... 152 9.3.6. Linear Extension Rate Measurements...... 152 9.3.7. Experimental Set-up...... 153 9.4. RESULTS ...... 154 9.4.1. OUR Measurements as affected by the Mass of Substrate and NaOH Pellets...... 154 9.4.2. OUR Readings as Affected by Substrate Moisture Content ...... 155 9.4.3. OUR Readings as Affected by Measuring Frequency...... 160 9.4.4. Mycelium Extension Rate...... 161 9.4.5. Yield and Substrate Bioefficiency as Affected by Substrate Moisture Content ...... 163 9.5. DISCUSSION ...... 164 9.6. CONCLUSIONS...... 167 CHAPTER 10 TWO ALTERNATIVE AGARICUS BISPORUS MUSHROOM PRODUCTION SYSTEMS USING GRAIN-BASED SUBSTRATES...... 168 10.1 ABSTRACT ...... 168 10.2. INTRODUCTION ...... 169 10.2.1. Steps in Agaricus bisporus Mushroom Production...... 172

viii 10.2.2. Factors Influencing Mushroom Yield in Grain-based Substrates...... 175 10.3. DESCRIPTION OF ALTERNATIVE MUSHROOM PRODUCTION SYSTEM ...... 176 10.3.1 Satellite Mushroom Production System ...... 176 10.3.2. Complete On-site Mushroom Production System ...... 177 10.3.3. Duration of Mushroom Production Process ...... 182 10.4. COST MODEL FOR GRAIN-BASED MUSHROOM PRODUCTION SYSTEM ...... 183 10.4.1. Cost Model for Satellite Mushroom Production System ...... 184 10.4.2. Cost Model for Complete On-site Mushroom Production System ...... 188 10.5. DISCUSSION ...... 192 10.6. CONCLUSIONS...... 194 CHAPTER 11 CONCLUSIONS AND SCOPE FOR FUTURE RESEARCH...... 195 REFERENCES ...... 200 APPENDIX A CULTIVATION OF PLEUROTUS ERYNGII ON SUBSTRATES COMPOSED OF GRAINS AND OILSEEDS...... 220 APPENDIX B RYE GRAIN SUBSTRATE FLOWABILTY...... 226 APPENDIX C COST MODEL EXCEL SPREADSHEETS...... 232 VITA......

ix List of Figures

Figure 2.1. Segmented-flow aseptic processing schematic (Walker and Beelman, 2002)...... 36 Figure 2.2. Continuous steam segmented-flow, aseptic processing system (Anderson and Walker, 2005) ...... 37 Figure 3.1. Effect of adding a single layer of perlite below the commercial grain spawn substrate containing 10% S41 on mushroom production ...... 49 Figure 4.1. Summary of the main effects (S41 rate, perlite volume, and casing type) on mushroom yield (kg/m2)...... 63 Figure 4.2. Summary of the main effects (S41 rate, perlite volume, and casing type) on BE (%)...... 64 Figure 4.3. Summary of significant interaction effects of S41 rate, perlite volume, and casing type on mushroom yield (kg/m2)...... 66 Figure 5.1. Mushroom yields for non-composted substrates composed of commercial millet grain spawn and different rates of five delayed-release supplements...... 77 Figure 5.2. Comparison of the temperature profiles during the 23-day period after casing for the production room (Env.) and non-composted substrates composed of commercial millet grain and five different five delayed-release supplements...... 80 Figure 6.1. Change in pH of peat moss heat-treated for different time intervals in an autoclave...... 93 Figure 6.2 pH for different casing treatments, before and after autoclaving...... 94 Figure 6.3. Comparison between activated carbon and NaOH as carbon dioxide absorbent...... 100 Figure 7.1. Environmental conditions and mushroom yield/day for A. bisporus tray bioreactor ...... 114 Figure 7.2. Environmental conditions and mushroom yield/day for A. blazei tray bioreactor ...... 115 Figure 7.3.Yield from two millet-grain substrates prepared on-site with one treatment receiving stage II supplementation, and compared to commercial grain spawn (Sylvan) substrate with stage II supplementation...... 116 Figure 7.4. A. bisporus fruiting on a substrate composed of grain and oilseed...... 118 Figure 7.5. A. blazei mushroom fruiting on a grain/oilseed substrate ...... 121 Figure 8.1. OUR measurements for A. bisporus and S. thermophilum at 16, 24, and 32oC...... 140 Figure 9.1. OUR observations as affected by the mass of the substrate with the red-line indicating OUR when all the observations are average...... 155 Figure 9.2. OUR of A. bisporus growing on a rye grain substrate with different substrate moisture contents treatments with rye grain spawn...... 157 Figure 9.3. Different moisture level treatments for a rye grain-based substrate inside the Oxtop® bottles...... 158 Figure 9.4. Change in porosity for the different rye grain substrate at different moisture levels at the beginning and at the end of the OUR measurement that spanned 32 days...... 159

x Figure 9.5. Mushroom yield and substrate bioefficiency for rye grain substrates from the Oxitop® bottles with different moisture levels supplemented with 14 g of Promycel Target®...... 160 Figure 9.6. Hyphal extension rate and lag phase duration for the different rye grain substrate with different moisture contents...... 162 Figure 9.7. Hyphal growth of A. bisporus on the rye grain substrate with different moisture contents...... 163 Figure 9.8. Mushroom yield and substrate bioefficiency for different grain-based substrates with the grain portion varying in moisture content...... 164 Figure 10.1. Overview of steps in Agaricus sp. mushroom production beyond the substrate preparation phase...... 174 Figure 10.2. A tray with mushroom fruiting from a substrate composed of millet grains and cracked soybean supplemented with Promycel Target® (delayed-release supplement) with an underlying layer of perlite...... 176 Figure 10.3. Overview of steps in the SMPS for Agaricus sp. mushroom production using a substrate of commercial grain spawn and delayed-release supplements...... 177 Figure 10.4. Four stages of the aseptic processing unit for non-composted grain-based substrate in the COMPS...... 180 Figure 10.5. Overview of steps COMPS for Agaricus sp. mushroom production using a substrate of cereal grains/oilseed substrate and delayed-release supplements...... 182 Figure 10.6 Breakdown of costs in % of total costs for the SMPS for the substrate requirements, tray requirements, labor and other costs (steam, electricity, and water) ...... 186 Figure 10.7. Change of mushroom production cost ($/kg) when yield per unit area (kg/m2) is increased for the SMPS...... 187 Figure 10.8. Change in mushroom price ($/kg) with respect to the change in price for commercial grain spawn ($/kg) for the SMPS...... 188 Figure 10.9. Breakdown of costs in % of total costs for the COMPS for the substrate requirements, tray requirements, labor and other costs (steam, electricity, and water) ...... 190 Figure 10.10. Change of mushroom production cost ($/kg) when yield per unit area (kg/m2) is increased for the COMPS...... 191 Figure 10.11. Change in mushroom price ($/kg) with respect to the change in price for millet grains ($/kg) for the COMPS...... 192 Figure A1. Pleurotus eryngii fruiting on oat-based substrates ...... 224 Figure B1. Mohr Coulomb yield criterion ...... 228

xi List of Tables

Table 2.1. Commercially available nutrient supplements for Agaricus bisporus...... 9 Table 2.2. Microorganisms prevalent in Phase II compost (Schisler, 1982) ...... 11 Table 2.3. Grain and oilseed nutritional quality based on percent wet weight basis (Morrison, 1959; Pomeranz, 1987; Nwokolo, 1996; Stamets, 2000, Weiss, 2000) . 26 Table 2.4. Physical properties of grains and oilseeds (Pomeranz, 1987)...... 26 Table 3.1. Mushroom yield from two different casing treatments, with and without AC 46 Table 3.2. Mushroom yield from grain spawn substrates treated with thiophanate-methyl and supplemented with S41 and S44...... 47 Table 3.3. Mushroom yield from grain spawn substrates without thiophanate-methyl application supplemented with S41 and S44 ...... 48 Table 4.1. Effect of different water-holding materials used as a single layer substratum on mushroom yield and bioefficiency...... 61 Table 4.2. Effect of adding perlite underneath a non-composted substrate of commercial grain spawn supplemented with S41 and cased with a sterilized casing containing 25% activated carbon or non-sterile casing on mushroom yield and substrate bioefficiency ...... 62 Table 4.3. Comparison between millet grain spawn and rye grain spawn on mushroom yield and substrate bioefficiency ...... 67 Table 5.1. Summary of yield, substrate bioefficiency and mean mushroom weight for a basal substrate composed of commercial grain spawn supplemented with S41, S44, Promycel Target, T6 and T7 delayed-release supplements...... 78 Table 6.1. Summary of treatments for both commercial grain spawn-based substrate and compost-based substrate...... 92 Table 6.2. Summary of results for casing treatments with a grain spawn-based substrate supplemented with 5% S41...... 96 Table 6.3. Summary of results for casing treatments with compost-based mushroom substrate supplemented with 5% S41 ...... 97 Table 7.1 provides a summary of the treatments applied for both fungi...... 112 Table 7.2. Summary of duration of the different steps in the mushroom production process...... 113 Table 7.3. Cultivation of Agaricus bisporus on different formulations of grain-based substrates...... 117 Table 7.4. Cultivation of A. blazei on different formulations of grain-based substrates120 Table 8.1. Treatments with different high temperature incubation periods with and without supplementation with Promycel Target®...... 133 Table 8.2. Treatments with shorter high temperature incubation periods all supplemented with delayed-release supplement Promycel Target® ...... 134 Table 8.3. Preliminary treatments for incubation of grain-based substrate with S. thermophilum ...... 137 Table 8.4. Colonization phase duration as influenced by the addition and pre-colonization of S. thermophilum in grain-based substrates ...... 138 Table 8.5. Incubation duration and grain type and their influence on mushroom yield, substrate bioefficiency, and average mushroom size...... 139 Table 9.1. Substrate formulations for the different moisture treatments ...... 149

xii Table 9.2. Observed OURs for treatments in which substrate mass and NaOH mass were varied...... 154 Table 9.3. Peak OUR for A. bisporus growing on a rye grain substrate with different moisture contents...... 156 Table 9.4. OUR of A. bisporus influenced by frequency of OUR measurements growing on a rye grain substrate with different moisture contents...... 161 Table 10.1. Description of individual processes occurring in the continuous aseptic processing unit for A. bisporus production on non-composted grain-based substrates in the COMPS...... 178 Table 10.2. Process durations for traditional compost-based system compared to the SMPS and COMPS...... 183 Table 10.3. Common parameters for the SMPS and COMPS cost models...... 184 Table 10.4 Parameters used for the energy calculations of the SMPS...... 185 Table 10.5. Parameters used for the energy calculations of the COMPS...... 189 Table B1. Failure strength of grain substrate at different confining pressures for the various moisture levels...... 229 Table B2. Summary of parameters of the mohr coulomb model...... 230

xiii Acknowledgments

The past five years at Penn State have been the most rewarding years in my life. I have grown both professionally and personally learning from my day to day experiences

as a graduate student. I would like to extend my sincerest thanks to Dr Paul Heinemann,

my thesis advisor, for his continued support over the years and for allowing me ample

freedom to pursue and develop my research. I would also like to thank Dr Paul Walker

for always having time to sit down and discuss my research. I really believe that these

talks have helped me tremendously in seeing the light at the end of the tunnel. I would

also like to thank Dr Pete Romaine for his time and effort to sit down with me and

discuss the mushroom side of things. Dr Romaine’s advice and direction were also

invaluable. I would also like to thank Drs Tom Richard and John Reagan for serving on my committee. I would like to thank Dr Richard for allowing me access to his lab, and

for always keeping me on track as to what engineering is all about. A special thanks is also in order for Dr Ali Demirci for his support and kindness. I would also like to thank the ABE department specifically Dr Roy Young for offering me the financial support to complete my graduate studies at Penn State.

I would like to extend a very warm thank you to all the faculty and staff members at the ABE department, Mushroom Spawn Lab, Mushroom Test and Demonstration

Facility, and the Mushroom Research Center. In particular, I would like to thank Dr

Virenda Puri, Vija Wilkinson, Tom Rhodes, Doug Keith, Henry Shawley, and John

Peccia.

Furthermore, I would like to thank all of my friends here in State College and elsewhere in the world. You know who you are…..whether it was tea’s at Café or beers at

xiv Otto’s, Mad Mex, Tony’s etc etc- all of it has been great. I extend a special thank you to

the Toney’s especially Mrs Evelyn Toney for their continued generosity and love

throughout these years and to Wren, Aalok, Maya, Art, Katherine, Dawn, Daniel and

Kadir for always lending an ear.

Finally, I would like to express my deepest gratitude to my parents who

throughout the years have supported and loved me. Without them all of this would not be possible. I would also like to thank my brother and sister for their love and for always being there to help me out and talk. I’ve missed you so much over the years. Last but not least, Walid what can I say….except that your friendship and love have shaped what I am today.

xv To my grandparents

Edmond and Hedda Naim

xvi

1. Introduction

The Agaricus bisporus mushroom is widely cultivated around the world, and

annual sales in the United States total 880 million dollars (USDA, National Agricultural

Statistics Service, 2005-2006). The conventional system of Agaricus mushroom

production relies on composting to produce a mushroom-specific substrate (compost

conditioned for mushroom growth is called substrate). Composting, the degradation of organic matter (animal and plant) into a soil-like material by microorganisms, is often a malodorous, lengthy (may take up to three weeks), and labor-intensive step in Agaricus

mushroom production (Derikx et al., 1990). Problems not only arise during the

production of the compost-based substrate, but also after the substrate is no longer

productive (known as spent mushroom compost or SMC) and is ready for disposal. Some

of the tribulations associated with SMC are odor production, salinity, and disposal methods. Even though mushroom production is no longer economically viable, SMC is still rich in nutrients that promote the growth of microorganisms associated with odor production (Heinemann et al., 2003). In fact, mushroom substrate is not fully composted, and this is why biological degradation of organic matter (composting) continues and odor production resumes. The other issue of concern when dealing with SMC is the limited venues by which mushroom producers may dispose of it. Disposal by land application is limited by the fact that SMC is rich in salts and nutrients that are capable of impairing the quality of surface water and groundwater (Williams et al., 2001; Guo et al., 2001). SMC is disposed by transforming it into a marketed soil amendment. However, it must be first composted completely before it can be sold as a finished product. This process may take

1 between nine months and 3 years if allowed to proceed passively (Guo et al., 2001;

Heinemann et al., 2003). SMC processing can be reduced to 10-12 weeks if the piles are

periodically turned. Costs involved, however, are high. As a result, an increased interest

in developing alternative mushroom production systems that are devoid of composting

has emerged. Most of the work until now has focused on the refinement of non-

composted substrate formulations composed primarily of agricultural and lignocellulosic

waste products. Only a handful of work has evaluated the use of grain-based substrates

for mushroom production.

Using grains as a substrate for the growth of the vegetative mycelium of the

mushroom fungus is a mainstay of the spawn industry (vehicle used to inoculate composted substrates with A. bisporus). Currently in the mushroom industry, batch

sterilization of grains using large V-blenders is used by a few spawn manufacturers for

the preparation of mushroom spawn, and for the most part the formulations and

conditions of growth are proprietary. Hence, there is little work in the literature that addresses the growth of A. bisporus on grains as opposed to compost.

Studies have shown that A. bisporus production is not limited to a substrate derived from composting (Till, 1962; Sanchez and Royse, 2001; Mamiro et al., 2007).

Production is also possible on various synthetic (non-composted) substrates, some of which are composed primarily of grain (Bechara et al., 2004). Benefits of using a grain- based substrate in mushroom production other than elimination of odor production include the following:

ƒ concentrated nutrient source that would reduce processing volume and waste

generation

2 ƒ more filling capacity in production chambers since substrate is shallow

ƒ short processing time (few hours)

ƒ spent grain substrate can be used as animal feed or as a bioenergy feedstock.

The overall research goal of this study was to develop a theoretical design and

refine material components of a grain-based mushroom production system (primarily for

A. bisporus) as a potential alternative to the environmentally problematic composting

system, and test whether this system could be used for the production of different types of

mushroom producing species. Specific objectives for this study were the following:

Objective 1

Perform preliminary test to determine the factors that significantly impact A. bisporus

mushroom production in grain-based substrates.

Objective 2

Define the factor level ranges that maximize A. bisporus mushroom production for

commercial grain spawn substrates.

Objective 3

Refine substrate formulations for A. bisporus composed of commercial grain spawn supplemented with delayed-release nitrogen-rich supplements.

Objective 4

Refine formulations of cereal grain and oilseed substrates supplemented with delayed-

release supplements and test whether different mushroom producing species (A. bisporus,

A. blazei, and Pleurotus eryngii) can be grown.

3 Objective 5

Determine whether co-cultivation with a mushroom compost thermophile (Scytalidium

thermophilum) would benefit A. bisporus mushroom production in grain-based substrates

and shorten spawn-run durations.

Objective 6

Test a range of substrate moisture contents for grain-based substrates and observe the

effects of these moisture levels on A. bisporus oxygen uptake rate, hyphal extension rate,

mushroom yield, and substrate bioefficiency with the goal of determining a targeted moisture level that maximizes mushroom production and shortens spawn-run duration.

Objective 7

Design the theoretical framework for the potential alternative grain-based system for A.

bisporus mushroom production including a study on the flowability of rye grain substrate

and some basic costs for the process.

Finally, this dissertation is presented as a paper-based format. Each individual

chapter can be treated as a stand-alone article or publication. This format was selected to

aid in the publication process.

4 2. Literature Review

Three subjects are covered within this section. One focus will be on A. bisporus mushroom production. Topics such as spawn production, nutrient supplements, traditional substrate preparation, and cultivation parameters will be covered thoroughly.

Since the growth of the mushroom fungus will be on solid substrates, second focus will be on solid-state fermentation (SSF). The two topics covered in this focus will be the various factors influencing fungal growth in SSF and the methods of fungal biomass quantification in such systems. The third focus will be on aseptic processing of particulate matter. This is necessary for the design of an aseptic processing system for grain sterilization. The combination of all these topics provides a suitable background for the design and implementation of a novel non-composted grain based mushroom production system.

2.1. Agaricus bisporus Mushroom Fungus

The nomenclature of the button mushroom was initially Psalliota hortensis

Lange but was renamed Agaricus bisporus Lange Sing in 1954 at the International

Botanical Congress in Paris. The button mushroom pertains to the group of

basidiomycetes and its life-cycle consists of four morphologically distinct stages: spore

formation, growth of hyphae which form mycelia (vegetative), primordia development,

and finally, maturation of the basidium, or mushroom. The basidium is composed of the

stipe or stalk and the pileus or cap. Spores are formed in the basidium to complete the life

cycle of the mushroom. There are three general A. bisporus mushroom strains: white, off-

white, and hybrid strains.

5 2.2. Spawn Production

Mushroom spawn is the vehicle by which the substrate is uniformly inoculated

with the mushroom fungus. The spawn medium is impregnated with a pure culture of

fungal mycelium and is generally composed of a solid organic matrix such as grains (rye,

millet, or wheat) or a liquid broth (Friel and McLoughlin, 2000). Initially, growers used spawn grown on a substrate primarily made of sterilized horse manure or tobacco stems.

Sinden (1932) developed the procedure to produce grain spawn, which involved placing

grains and water in a cotton-wool plug sealed bottle, sterilizing it with steam for more

than an hour, and finally inoculating the sterilized grains with a pure culture of Agaricus.

Guiochon (1958) applied for a patent in which he described a method of producing spawn

in heat-resistant bags. Stoller (1962) described a grain spawn preparation method in

which 10 kg of wheat grain and 15 L of water were boiled for 15 minutes after which

they were allowed to soak for an additional 15 minutes. The water was then removed and

the wheat grains were allowed to dry. When the grains cooled, 9 kg of cooked grains

were mixed with 120 g of gypsum and 30 g of calcium carbonate. The purpose of the

gypsum was to prevent the grains from sticking, whereas the calcium carbonate was used

to adjust the pH (Stoller, 1962). Lemke (1971) developed what is termed as “perlite

spawn”. Perlite, a heat-expanded inert mineral, was impregnated with a nutrient solution,

filled in bottles, sterilized, and finally inoculated with Agaricus mycelium. The “perlite

spawn” substrate was composed of the following ingredients: 1450 g perlite, 1650 g

wheat bran, 200 g gypsum, 50 g calcium carbonate, 6650 ml of tap water. Stoller (1972)

describes the production of finely ground mushroom spawn grown on a substrate of

grain, soybean meal, casein, and yeast.

6 Romaine and Schlagnhaufer (1992) developed an alginate-based mushroom spawn. Pellets were composed of calcium alginate, vermiculite, hygramer, and nutrisoy

(as nutrient source). When used as inoculum for composted substrate, the performance of the alginate-based spawn was comparable to grain spawn.

Kananen et al. (2000) developed and patented a spawn medium (to be used as inoculum for commercial composted substrate) which was termed as mushroom-spawn supplement. They formulated several grain and supplement mixtures (commercial and non-commercial), inoculated the mixtures with the mushroom fungus, and used the colonized spawn medium to inoculate a compost-derived substrate. The average yield of mushrooms from a compost-based substrate inoculated with the spawn-supplement mixture was greater than a similar substrate inoculated with the traditional rye grain spawn. Apart from greater mushroom yield, Kananen et al. (2000) also reported lower incidence of disease in the substrates inoculated with mushroom spawn-supplement and shorter spawn runs (rapid colonization of the composted substrate).

The moisture content of mushroom grain spawn should range between 38-55%.

A moisture content below this range hinders the mycelium colonization process and anything greater than 55% may promote the growth of bacteria (Stamets, 2000a).

Most spawn is commercially produced by various spawn suppliers (Sylvan

Spawn Laboratory; Amycel, L. F. Lambert Spawn Co.). Sylvan uses large V-blenders

(double cone blenders) in which the solid nutrients (grains and other materials) are mixed, sterilized, and inoculated with Agaricus mycelium. The inoculated material is aseptically added to sterile bags with breathing patches and then transferred to environmentally controlled rooms where the mushroom fungus colonizes the solid

7 matrix. Finally, the finished grain spawn is stored in refrigerated chambers and transported to customers.

In the Mushroom Spawn Laboratory at Pennsylvania State University, Agaricus grain spawn is produced predominantly on rye grain (200 g rye grain + 4 g of calcium carbonate+ 4 g of calcium sulfate + 220 ml water). The grain mixture is placed in 1000 ml flasks capped with cotton wool plugs and autoclaved for 45 min at 121°C (fast exhaust).

2.3. Nutrient Supplements Used by Mushroom Producers

Studies have shown that Agaricus mushroom yield can be increased by adding various supplements to commercial mushroom substrate. Carroll and Schisler (1974) describe using denatured protein as a nutrient supplement in compost-based substrates.

The protein was mixed into the substrate before or shortly after spawning. In addition,

Carroll and Schisler (1976) showed that encapsulating micro-droplets of vegetable oil within a protein coat denatured by formaldehyde increased mushroom yield on compost.

Romaine and Marlowe (1993, 1995) developed an intact seed-based, delayed-release nutrient supplement for Agaricus grown on compost. One type of seed they investigated was canola (rapeseed). The canola embryo was inactivated using heat treatment (drying oven at 95 °C for 24 hrs or autoclaving at 121 °C for 1.5 hrs) to prevent germination and was added to compost. Mushroom yield from the canola-supplemented compost was comparable or greater than mushroom yield from compost supplemented with

Spawnmate II SE (a commercially available supplement).

Today, various suppliers offer several mushroom nutrient supplements

(relatively high in nitrogen) with delayed-release and antifungal properties. Table 2.1

8 provides a summary of some of the mushroom nutrient supplements available on the

market.

Table 2.1. Commercially available nutrient supplements for Agaricus bisporus. Supplement Supplier %Nitrogen % Protein % Fat % Fiber S41 Full House 5.75 36 18.75 5.7 S44 Full House 6.25 40 18.75 5.70 T5 Full House 8.80 55 13.15 5.80 T6 Full House 6.75 42 3.50 30.75 T7 Full House 10.55 66 2.25 14.75 Promycel Amycel NA 60 NA NA 600 X-cell 55 Sylvan NA 54-56 3-4 NA CS36 Sylvan NA 34-36 16-18 NA Milli Champ Sylvan NA 46-48 3-4 NA SpawnMate Amycel NA 27 NA NA II SE

2.4. Commercial Production of Agaricus bisporus

Agaricus bisporus, as are all fungi, is a heterotrophic organism; it lacks

chlorophyll and cannot use photosynthesis to supply its carbon source. Carbon must be

supplied from organic matter. In commercial settings, the mushroom fungus nutrients are

derived from a partially composted mixture of animal and plant organic matter.

2.4.1. Composting in Commercial Agaricus bisporus Production

The mushroom industry prepares a suitable substrate for Agaricus mushroom production by composting plant and animal organic matter. The result is the production of a substrate that provides sufficient nutrients to promote mushroom production and inhibits the growth of competitor organisms. Generally, there are three methods of composting: long-term method, short term, and indoor composting (Rai, 2004). The long- term method, the oldest, is still adopted in certain parts of the world and consists of composting the organic material outdoors for 28 days. This method, highly inefficient, is

9 prone to contamination from competitor microorganisms (Vijay and Gupta, 1995). The short-term method of composting was developed by Sinden and Hauser (1950) and consists of two phases (San Antonio, 1975; Schisler, 1982), Phase I and Phase II. In

Phase I the microorganisms start metabolizing the organic matter by breaking it down into simpler forms. Mixing and periodic watering of the piles promote the uniform breakdown of the organic material. As the microorganisms start increasing in number, the temperature of the compost piles starts increasing. This temperature increase provides a suitable environment for the growth of mesophilic and thermophilic microorganisms.

Much of the nitrogen is then ammonified (Schisler, 1982). When the temperature of the compost piles reaches 80 ºC, most of the activities turn to chemical and almost all biological activities cease. The major chemical reactions taking place are the carmelization of carbohydrates and browning, or Maillard reactions. Carmelization of carbohydrates produces dark colored polymers as a result of consecutive dehydration and condensation (Schisler, 1982). Maillard reactions, occur at elevated temperatures when amino acids, peptides, or proteins react with hemiacetal hydroxyl groups of sugars. Phase

I is considered complete once the material is dark brown in color with a sharp odor of ammonia and once the material becomes pliable and water absorbent. The material is then ready for Phase II.

The composted material from Phase I is transferred to Phase II “tunnels”. The steps involved in this composting stage are twofold. First, the composted material is pasteurized by maintaining a temperature of 60 °C for 2 hrs. Following pasteurization, fresh air is introduced into the tunnels in order to reduce the compost temperature to 50

°C and eliminate ammonia (Schisler, 1982). Maintaining temperatures of 50 °C for 4-6

10 days provides a suitable environment for the growth of mesophilic and thermophilic microorganisms. As a result of this growth, ammonia and other compounds are assimilated into the microbial biomass which at later stage are used as a source of nutrients for the growth of A. bisporus (Wood and Fermor, 1985; Ross and Harris, 1983).

In fact, Ross and Harris (1983) attribute the selective nature of compost for A. bisporus to the growth of these thermophilic and mesophilic microorganisms. They showed that inhibiting the growth of mesophiles/thermophiles, either by excessive heating or through the addition of volatile antimicrobials such as chloroform and alcohol, compost no longer was selective for A. bisporus. However, selectivity could be restored after inoculating non-selective compost with a thermophilic fungus Torula thermophila or fresh compost and re-subjecting the material to Phase II composting. Their interpretation was that an intact (non-lysed) but dormant thermophilic/mesophilic biomass is conducive to compost selectivity. Table 2 provides a few examples of mesophilic and thermophilic organisms found in compost during Phase II.

Table 2.2. Microorganisms prevalent in Phase II compost (Schisler, 1982) Microorganisms Thermomonspora, Streptomyces, and Actinomycetes Thermoactimomyces Tortula, Chaetomium, Mucor, Scytalidium Fungi thermophilum and Apergillus fumigatus

The indoor composting came about as a result of environmental problems related to composting; odors were deemed the most problematic. Laborde (1992) described indoor composting which was categorized into two methods: INRA (Institut Nationale de la Recherche Agronomique) method and the Anglo-Dutch method. The difference between these two methods was mainly temperature. For the INRA method, Phase I was

11 carried out at 80 °C for 2-3 days followed by Phase II at 50 ºC for 5-7 days (Laborde,

1991). The Anglo-Dutch method entailed conditioning the substrate at 41 ºC for a week

followed by a short pasteurization phase at 60 ºC for 4-6 hrs.

2.4.2. Mushroom Fungus Spawning

A. bisporus spawn is used to inoculate the substrate with Agaricus mycelium.

Spawning is the phase during which mushroom growers incorporate the A. bisporus

spawn into the substrate. The mycelia, the vegetative stage of the mushroom fungus, in turn, colonize the substrate by utilizing the available nutrients within the substrate. The

optimal economically viable rate of grain spawn application is around 400-600 g/m2 of bed area containing 70-80 kg of substrate (Flegg et al., 1985). However, this rate changes depending on the price of grain spawn. Grain spawn generally gives better results compared to manure spawn, and increasing the rate of spawning increases yield (Flegg et al., 1985). At Penn State’s Mushroom Test Demonstration Facility, the amount of

Agaricus grain spawn applied is 2.5% of the weight of the substrate. To begin the spawning phase, the composted substrate that comes out of Phase II is transferred to mushroom growing units such as wooden trays. Some studies have shown that growing

Agaricus in polythene bags is practical, and a 25 cm deep compost-based substrate produced the greatest yield of 13.9 kg/100 kg of compost in a 50 day cropping interval

(Gupta et al., 2004). After the spawn is incorporated into the substrate, the inoculated substrate trays or beds are kept at 22 ºC in a non-ventilated room with a relative humidity of 75%-80%. A. bisporus mycelia start colonizing the substrate.

12 2.4.3 Substrate Casing and Casing Inoculum

Once the mycelium covers the substrate surface (10-14 days) and 50% or more of

the surface is colonized, the substrate is cased (casing being a mixture of peat moss and

calcium carbonate). The casing layer is spread evenly over the colonized substrate. It serves as a mechanical support, water, and microflora (indigenous to the peat moss) for

the development of mushrooms. Flegg (1956) defined the role of the casing as a layer “to induce sporophores (mushroom pin heads) in quantity”. There are different types of casing layers; a mixture of peat moss and lime is currently the most common. The lime will bring the pH of the peat moss up to 7-7.5 which is ideal for mushroom production.

Once the substrate is colonized (50% or more) with A. bisporus mycelia, the colonized substrate is covered with a casing layer. The casing may also be applied right after spawning, although the casing has to be watered with particular care to avoid the formation of a crust (Flegg et al., 1985). After covering the substrate with the casing layer, the mushroom fungus is pinned right after the mycelia emerge on the top of the casing layer and cover 50% or more of the surface (7-9 days).

In many commercial operations, casing inoculum (CI) is added to the wetted casing and then spread over the entire surface of the substrate. Casing inoculum is composed in part of an inert carrier material (vermiculite) and a nutrient source such as bran. The material is sterilized and then inoculated with the mushroom fungus. Once fully colonized the material is added to mushroom casing. The addition of CI shortens the production time and improves the quality and yield of mushrooms (Gupta et al., 1989).

Many theories have been presented explaining what exactly triggers the formation of pins in the casing layer. Tschierpe (1959) concluded that the role of the casing layer

13 was to create a carbon dioxide gradient and this gradient would trigger the fruiting of

Agaricus mushroom. Eger (1961) showed that microorganisms present in the casing layer are responsible for the fruiting of A. bisporus. The number of plausible theories may have driven Peerally (1981) to state that the most elusive process of A. bisporus is the fruiting.

He concluded from his studies that A. bisporus failed to fruit on agar if not treated with a solution of casing bacteria. Hume and Hayes (1972) claimed that cultures of

Pseudomanas putida, which promote fruiting of Agaricus in compost, stimulated the formation of pins with Agaricus cultures grown on sterile malt extract agar. This observation was also repeated by Long and Jacobs (1974). However, they also showed that activated carbon incorporated into a sterile casing layer at 25% (w/w), was able to promote the formation of mushrooms as well. The activated carbon used in their experiments was specific for the adsorption of gases. Verbeke and Overstyns (1991) showed that activated charcoal influenced reactions of carbon dioxide in water resulting in reduced calcium. Oxalate, which has iron chelating properties, is secreted by the mushroom mycelium. When iron chelation via oxalate occurs, the iron is more prone to be reduced. The reduced form of iron is believed to inhibit the formation of pins. Calcium reduces the solubility of oxalate thus reducing the amount of reduced iron. However, when the casing is sterilized, the heating process interferes with the liberation of calcium, which in turn interferes with the precipitation of oxalate and ultimately reduces the yield of mushroom through the reduction of iron. In summary, the exact role of the casing layer in the mushroom production process is still unknown (Romaine and Schlagnhaufer,

1992).

14 2.4.4. Mushroom Fungus Pinning and Harvesting

Environmental conditions in the mushroom production chambers are adjusted in order to promote the fruiting of A. bisporus. Reducing the temperature to 16 ºC and increasing ventilation triggers fruiting. The ventilation should reduce the carbon dioxide level from 5000 ppm to about 800 ppm (Tschierpe, 1959). This stage is called “pinning”, because it is the phase during which the mushroom fungus is triggered to form pins, the precursors of mushroom fruit bodies. The pins increase in size to develop into harvestable mushrooms, which takes around 7-10 days from initiation of pinning.

Mushrooms are harvested by hand. There has been work on developing harvest machines that would save on labor (Persson, 1972), although few harvest machines, if any, are used commercially. Generally, mushrooms are produced in a series of harvests also known as “breaks” or “flushes”. The duration of harvest for each flush can take up to

3-6 days while the average time between each flush is around 6-12 days (Flegg, 1956).

The first flush will produce about 40%-50% of the total mushroom yield. Although mushrooms will continue to be produced for several weeks, mushroom growers will cease harvesting when it is no longer economically viable (usually 3-4 weeks). For 2002-

2003 the average yield of mushrooms for the U.S. was 6.3 lb/ft2 or 31 kg/m2.

2.4.5. Metabolism and Physiology of A. bisporus

A. bisporus, as all fungi, produces extracellular enzymes that degrade various compounds. The degraded nutrients are then absorbed into the mycelium to be used for growth, maintenance, and mushroom production. In composted substrates, the enzymes released by A. bisporus are: laccase, endocellulase, xylanase, proteases, lipases, phosphatase, and laminarinase (Wood and Fermor, 1981). Laccase degrades lignin and

15 the phenolic fraction of the substrate, whereas laminarinase degrades the glucan

(carbohydrate) fraction. Studies show that the nutrients consumed by the mushroom fungus are dependent on the stage of growth. During the vegetative stage, the

assimilation of the lignin fraction is correlated with an increase in laccase production

(Wood and Fermor, 1985). Gerrits (1988) showed that 63-92% of the lignin fraction is

consumed from spawning until pin formation. However, during the fruiting stage,

cellulose and hemicellulose are the major fractions which are consumed. During this

stage, a decrease in laccase production and increase in endocellulase release is observed.

The carbohydrate metabolism within the mushroom fungus also undergoes changes

during the vegetative and fruiting stages. Glycogen and trehalose are accumulated in the

mycelia during vegetative growth and are thought to be storage carbohydrates which are

later used for mushroom production (Wannet et al., 1995). Manitol, a sugar alcohol, is

accumulated in the fruiting bodies which is thought to act as an osmoticum during growth

and then as a respiratory substrate after harvest (Hammond and Wood, 1985; Wannet et

al., 1995).

The role of microbial biomass (living and non-living) as a nutrient source for A.

bisporus has also been investigated (Wood and Fermor, 1985). During substrate

preparation (Phase I and Phase II composting), microorganisms grow on the organic

material forming a polysaccharide matrix embedded with microbial cells composed of

40% carbohydrates, 12% protein, and 4% phenolic compounds (Wood and Fermor,

1985). This polysaccharide matrix contains a large percentage of compost nitrogen.

Laboratory studies have also shown that A. bisporus is capable of degrading heat-killed

bacterial cells and uses them as sole carbon and nitrogen sources. In addition, studies

16 have shown that A. bisporus is capable of degrading various thermophilic fungi isolated

from compost such as Humicol insolens and Pennicilium chrysogenum, among others

(Wood and Fermor, 1985). One of the compounds entrapped within the thermophilic

biomass is linoleic acid, which is the predominant fatty acid. Linoleic acid stimulates

mushroom growth in composted substrates but not in liquid cultures (Dijkstra et al.,

1972; Schisler, 1982). Interestingly, linoleic acid is the precursor of the key component of the characteristic mushroom flavor 1-octen 3-ol (Morawicki et al., 2005).

The mechanism of translocation of material within fungi is not well understood

(Hammond and Wood, 1985). Diffusion, osmotic flow, and transpiration have all been

proposed (Amir et al., 1995). However, the mechanism of translocation in mushrooms is

thought to be largely due to bulk flow induced by evaporation (Hammond and Wood,

1985). Translocation of radio-labeled glucose within the mycelia of Armillaria mellea

was found to occur only under aerobic conditions, whereas absorption with no

translocation occurred under anaerobic conditions (Anderson and Ullrich, 1981). The

mechanism of flushing, production of a series of sequential mushroom harvests, is hypothesized to be due to the accumulation of products within the mycelium (glycogen and trehalose) which trigger the formation of mushrooms. Once pins start forming, these products are reduced below a threshold level preventing the formation of new mushrooms

(Hammond and Wood, 1985). Threshold levels are restored after mushrooms are harvested.

Minerals are required by various fungi for growth. Magnesium, calcium, iron, copper, manganese, zinc are some of the essential minerals for fungal growth (Jennings,

1995). The concentration of calcium in the growing medium required by fungi is among

17 the highest of the previously stated elements (Royse and Sachez-Vasquez, 2003).

Calcium ions are concentrated in the apical regions of growing tips of hyphae. It is

thought that calcium ions enter the hyphae through passive mechanisms and are expelled by energy-dependent mechanisms. The main role of calcium is in regulation of hyphal apices and the formation of hyphal branches (Gadd, 1995). In addition, a high calcium

gradient within hyphae was shown to promote the elongation of hyphal apices of

Neurospora crassa (Maheshwari, 2005).

Mushrooms consist of 90% water. This water is obtained from the substrate and

casing layer and is a factor that influences the growth and yield of mycelia and

mushrooms, respectively (Flegg, 1985). Increasing the amount of water applied to the casing layer is correlated with an increase in mushroom yield (Flegg, 1985). In composted substrates, the best growth of vegetative mycelium was observed for substrates with moisture contents about 55% to 70% (Flegg et al., 1985). Wetter substrates packed less densely were characterized with better growth and this was assumed to be due to better aeration (Flegg et al., 1985). An empirical relationship which defines the water balance within the substrate and casing is shown in equation 1 (Flegg,

1985).

Q = 1.8(1.25P + E + K − 0.1CDM − 0.18TW ) Eq. 1

where Q = total water required

P = fresh weight of the crop harvested

E= surface evaporation loss

K= cst = 280 kg/100 m2 bed

CDM= compost dry matter

18 TW= total water of the compost and casing at start

The constant K accounts for minor losses of water in bottom and sides of trays after watering. Another important characteristic of mycelium and its relation to water is the increasing hydrophobicity of the substrate (Flegg, 1985). For instance, excessive growth of mycelium on the casing layer renders it almost impermeable to water. Fungal mycelia secrete proteins termed as hydrophobins that are used to make hydrophilic surfaces hydrophobic and hydrophobic surfaces hydrophilic (Wosten and de Vocht, 2000). These proteins are involved in lowering surface tension by covering the emerging aerial hyphae and propagative structures such as mushrooms with a hydrophobic layer resulting in the growth into air from a liquid medium (Askolin et al., 2005).

2.5. Agaricus Mushroom Production on Non-composted Substrates

Till (1962) developed what is commonly known as the Till substrate, which consisted primarily of chopped and ground straw, white peat, calcium carbonate, cottonseed meal and soybean meal. After mixing, moistening, and sterilizing the substrate, it was inoculated with A. bisporus. The mycelium took longer to grow compared to normal compost. However, the yield in kg/ton of substrate was high. This process was never adopted because of high operating costs. Huhnke and Von Sengbush

(1968) simplified the Till process by replacing sterilization with pasteurization. The substrate was steamed and then inoculated with a suspension of microorganisms from compost. However, the results were inconsistent.

Smith and Hayes (1972) grew A. bisporus on inert sphagnum peat moss with nutrients supplied in a solid form. The maximum yield obtained was 1.04 kg/m2 which was higher than the liquid nutrient solution treatment (0.44 kg/m2).

19 A method of producing A. bisporus fruiting bodies from cased grain spawn was developed by San Antonio (1971). Rye grain mixed with calcium carbonate and de- ionized water (20 g rye grain/0.4 g CaCO3/20 ml di-water) was sterilized (121°C for 60 min). Following sterilization, the rye grains were inoculated with 10-20 grains of spawn.

Mushroom spawn was left to grow aseptically on the grains and once fully colonized, a

layer of pasteurized casing was placed on the colonized grains. When the mycelium

reached the surface, aeration was increased and fruiting bodies were obtained. Although

yield of mushrooms was not presented, San Antonio (1971) concluded that the quantity

of mushrooms produced was comparable to that obtained from conventional compost.

Sanchez and Royse (2001) showed that a pasteurized mixture of oak sawdust,

millet, rye, peat, alfalfa meal, soybean flower, wheat bran, and calcium carbonate was

suitable for the production of A. bisporus (brown Portobello). The maximum biological

efficiency (fresh weight of mushrooms divided by the dry weight of the substrate)

2 achieved was 77.1% and mushroom yield was 31.4 kg/m .

Bechara (2004) investigated different non-composted, grain-based substrates. The

substrates were primarily composed of grains mixed with a proportion of perlite (water-

holding material) at ratios of 100/0, 75/25. 50/50 and 25/75 (grain/perlite). The substrates were uncolonized millet mixed with various proportions of perlite, and commercial rye grain spawn (used as a substrate) mixed with various proportions of perlite. For the millet

substrate, the 75/25 treatment produced 7.69 kg/m2 and was comparable to compost- grown mushrooms which produced 8.04 kg/m2. The yield from the compost grown

mushrooms was lower than commercial yield (31 kg/m2) because in the former a 4 cm

layer of commercial substrate was used (comparable depth to the millet substrate),

20 whereas in the latter a 12-20 cm layer of commercial substrate is typically used. The

yield of the 100/0 commercial grain spawn to perlite treatment was the highest of the

grain spawn treatments, producing 5.3 kg/m2, while for the 75%, 50%, and 25%

commercial rye grain spawn treatments, yields were 3.9 kg/m2, 3.0 kg/m2 and 2.2 kg/m2,

respectively. Overall, treatments with millet grain were the most productive in terms of

mushroom yield.

Mamiro (2007) used a non-composted substrate composed of sawdust (28%),

millet (29%), rye (8%), peat (8%), ground alfalfa (4%), ground soybean meal (4%),

wheat bran (9%) with and without the addition of pasteurized SMC. The highest yield

and biological efficiency obtained were 27.2 kg/m2 – 144.3%, and they were observed for

treatments of 1:1 ratio of non-composted substrate and SMC spawned with casing

inoculum and supplemented with 10% Promycel Target® (Mamiro, 2007). Furthermore,

the addition of 0.9% Micromax, a mixture of minerals specifically used in mushroom

substrate, significantly increased mushroom yield from 8.5 to 14.6 kg/m2.

Sanchez et al., (2007) tested a non-composted substrate composed primarily of

Pangola grass and showed that pre-treating the substrate with S. thermophilum improved

substrate bioefficiency from 11 %, for the control group, to 26.4% for pre-treated substrate. The authors state that until now, there is not an economically-viable alternative

to the traditional mushroom production system, even considering a system using grain-

based substrates.

2.6. Agaricus bisporus Used in Animal Feed

Studies have explored the potential of using Agaricus bisporus mycelium as

animal feed. Torev (1968) indicated that mushroom mycelium contains about 45%

21 protein. When used as a bio-concentrate ranging from 3%-5% of total feed formulation

mass in pig, calf, and broiler feed, an 18% to 22% increase in growth was detected. For

laying hens, including mushroom mycelium in their feed increased egg production 14%

to 18%.

In addition, supplementing mushroom tissue to animal diets was shown to reduce

phosphorous pollution. Most cereals and legumes are rich in phytic acid (Liu et al.,

1998). Phytic acid is an anti-nutritional factor that reduces mineral absorption in

monogastric animals. Decreased phosphorous uptake from feed due to the presence of

phytic acid has lead to increased phosphorous pollution from manure in areas of intense

animal production (Common, 1989; Nasi, 1990). Several fungi produce phytase, an enzyme which hydrolyzes phytic acid. Phytase content of A. bisporus was shown to be

0.211 U/g of tissue (Collopy, 2004). Hence, using grains colonized by the mushroom

fungus as animal feed is a promising area, especially for its potential in increasing

phosphorous availability to animals.

2.7. Solid State Fermentation

The growth of A. bisporus on a compost-derived substrate or a cereal grain-based

substrate is a solid state fermentation (SSF) process. Hence, this section will introduce

the topic of SSF and the various process variables and methods of fungal biomass

estimation in SSF systems.

2.7.1. Definition of Solid State Fermentation

Solid-state fermentation is a microbial process occurring on solid or granular

substrates, either inert, or organic, used to support growth in the absence of free flowing

22 water (Nagel, et al., 2000; Ooijkass et al., 2000; Pandey et al., 2000). The term solid

substrate fermentation, as recommended by Pandey et al. (2000), is to be used whenever

the substrate is of an organic nature and the growing microorganism derives its nutrients

from it. Hence, from here on the term solid-state fermentation (SSF) will be used.

Currently, various products are derived from solid-state fermentation; food, bio-

pesticides, plant growth hormones, enzymes, feed, pharmaceuticals, and nutraceuticals

(Ooijkass et al., 2000; Pandey et al., 2000). Miller (1996) challenges the use of the term

solid-state fermentation. According to the author, the word solid is misleading and may

imply the material is devoid of pores which is not the case. Furthermore, the word

fermentation is typically used to describe an anaerobic process which again is not the

case since most SSF are aerobic. Therefore, Miller (1996) proposes a new term which is

matric phase culture. However, for the purpose of this body of work SSF will be adopted.

2.7.2. Support Material Used in SSF

Ooijkass et al. (2000) distinguish two types of support media in solid state

fermentation; inert and organic substrates. The inert substrate, not predominantly used in

SSF systems, is impregnated with a liquid nutrient medium from which the

microorganisms derive their nutrient source. Some examples of inert materials that have

been used are vermiculite for the cultivation of Colletotrichum truncatum, perlite for the

cultivation of A. bisporus, polyurethane for the cultivation of Aspergillus oryzae, and polystyrene for the production of Vibrio coticola (Ooijkass et al., 2000; Bechara et al.,

2004). Examples of organic support materials are: cereal grains for Aspergillus panasitus, bran for Gibberella fujikuroi, sweet potato residues for Streptomyces viridifaciens, compost for A. bisporus, cottonseed hulls for Pleurotus ostreatus, and various

23 ligninocellulosic materials (Ooijkass et al., 2000; Pandey et al., 2000). One of the main disadvantages of using organic supports is that the nutrients for microbial growth constitute part of the support structure. Hence, with degradation the geometry and physical characteristics of the material will change (Ooijkass et al., 2000). In addition, the recovery of products from inert supports is easier than from organic supports (Ooijkass et al., 2000). Finally, inert supports allow the use of chemically defined media for the growth of microorganisms which reduces heterogeneity and improves reproducibility.

Generally, organic substrates are products or by-products of agriculture and the agro-industry. The various components of each substrate are composed of cellulose, hemicellulose, lignin, pectin, proteins, fats and other compounds (Raimbault, 1998). In cereal grains, the major carbohydrate source is starch, which is composed of amylose

(16-30%) and amylopectin (65-85%). Both starch components are composed of monomers of glucose. However, starch will only become available to microorganisms after it has gelatinized (although there are some microorganisms which can degrade ungelatinized starch) (Raimbault, 1998). Gelatinization of starch will be discussed in a later section.

2.7.3. Grains and Oilseeds

The focus of this section is on the nutrient composition of the following cereal crops: rye, oats, and millet. For oilseeds, this section focuses on the nutrient composition of canola, niger, and soybean. However, nutritional composition of both cereal and oilseed crops vary widely because of differences in varieties and growing conditions.

24 Rye, Secale cereale L., contains 10% protein on a dry weight basis and is

characterized by having high levels of the amino acid lysine. Much of the rye in the US

is used as feed grain, whereas it is used as a bread grain in Europe.

Millet, Panicum miliaceum, is used for human consumption in many areas of the

world. It has also been found to be suitable as animal feed. It contains about 5% to 8%

oil. Although deficient in essential amino acids, it contains adequate levels of lysine

compared to other grains. It also contains 27% to 32% more protein than maize (Smith et

al., 1989).

Oat, Avena sativa, production used for human consumption and animal feed grain

has decreased over the years (Youngs and Forsberg, 1987). The oat hull is primarily

composed of cellulose and houses the highly nutritious groat. Oat grain is composed of

11-14% protein, 55-65% carbohydrate, and 4-7% fat (Stamets, 2000a; Youngs and

Forsberg, 1987).

Canola (Canada Oil Low Acid), is a trademark cultivar of rapeseed, Brassica

napus L. During the 1960s, research showed that common rapseed contained high levels of erucic acid which is mildly toxic to humans in large doses. However, Canada started a breeding program that produced rapeseed varieties with low erucic acid content (less than

2%). The oil content of canola is 38%.

Niger, Guizotia abyssinica, is an oilseed originating from the northern Ethiopian

highlands. The seed oil content may vary between 25% to 45%, whereas the protein

content may vary between 12% to 25% (Weiss, 2000).

Soybean, Glycine max, is used in various foods and feeds for humans and

animals, respectively. The US is one of the largest producers of soybean which is

25 characterized by having an oil content range of 15-22% and a protein content of 40-50%

(Weiss, 2000). The protein portion contains almost the entire complex of amino acids including the essential amino acids for human consumption (Weiss, 2000). Tables 2.1 and 2.2 summarize the nutritional components and physical properties of the grains and oilseeds.

Table 2.3. Grain and oilseed nutritional quality based on percent wet weight basis (Morrison, 1959; Pomeranz, 1987; Nwokolo, 1996; Stamets, 2000, Weiss, 2000) Protein Carbohydrate Lipid/Oil Ash Fiber Grain/Oilseed Content (%) (%) (%) (%) (%) Rye 12.6 70.9 1.7 2.1 2.4 Millet 11.9 63.7 3.4 3.4 8.1 Oats 12.0 58.6 4.6 4.0 11.0 Niger1 17.9 17.9 38.5 4.6 11.9 Canola 20.4 15.7 43.6 4.2 6.6 Soybean 37.9 24.5 18.0 24.5 5.0

Table 2.4. Physical properties of grains and oilseeds (Pomeranz, 1987). Bulk Length Width Grain Wt Grain/Oilseed Density (mm) (mm) (mg) (kg/m3) Rye 4.5-10 1.5-3.5 21.0 695 Millet 1.5 1.5 - - Oat 6-13 1.0-4.5 37.0 790-825 Soybean NA NA NA NA Canola NA NA NA NA Niger NA NA NA NA

2.7.4. Environmental and Process Parameters in SSF

Fungal growth in SSF is influenced by various environmental and process parameters such as temperature, pH, water activity, and aeration (Raimbault, 1998).

Sterility of substrates is also another important parameter in SSF.

Temperature control is important in any biological process. In solid-state fermentation it is critical to have a suitable control of temperature because overheating,

26 due to microbial activity along with reduced rates of heat transfer (as compared to

submerged fermentation), may hinder growth (Nagel et al., 2000). Evaporative cooling,

coupled with water spraying, was shown to be more effective than convective and

conductive cooling (Barstow et al., 1988; Nagel et al., 2000).

The pH of the solid substrate may change as a result of microbial activity. Certain

microorganisms, such as Aspergillys sp., tend to lower the pH far more than others

(Raimbault, 1998). In addition, the substrate used also influences pH kinetics, for

instance, lignocellulosic materials tend to have a buffering effect (Raimbault, 1998).

However, if the pH needs to be controlled, it is difficult to adjust it during the

fermentation process. Therefore, the addition of buffers such as calcium carbonate, as

used in the mushroom spawn industry, or various nutrients (urea and ammonium) can control the pH (Raimbault, 1998).

Water in solid substrate fermentation is found sorbed to the matrix and is usually

limiting (as compared to submerged fermentation). The moisture content of various solid

substrates may be comparable, however, their water activity (free water available for

microbial growth) may be different (Oriol et al., 1988). Hence, measuring water activity is preferred over measuring moisture content of solid substrate. Water activity is defined

by the following equation (Aw) = RH/100 = P/P0 where P is vapor pressure of water in

the substrate and P0 is vapor pressure of pure water at the corresponding temperature

(Griffin, 1981). Calibrated relative humidity sensors can be used to measure water

activities of solid substrates such as compost (Bloom and Richard, 2002). Optimal

moisture content for growth and substrate utilization ranges 40 to 70% (Raimbault,

1998). Optimal water activity depends on the microorganism. Bacteria require water

27 activities greater than 0.9, whereas fungi require water activities greater than 0.68 with

the optimal water activity ranging from 0.8 to 0.96 (Raimbault, 1998).

The role of aeration in SSF, other than maintaining aerobic conditions, is to regulate substrate temperature and moisture level (Raimbault, 1998). Convective air movement within solid substrates is rapid, allowing high rates of oxygen diffusion. The measurement of total air space within a solid matrix is air filled porosity. Air filled porosity is the volume fraction of air in a porous matrix that can be measured using a gas pycnometer (Richard et al., 2002). The equation for air filled porosity is the following:

Vg εa = Eq. 1 Vg + Vw + Vs where Vg, Vw, and Vs are the volumes of gas, water, and solids in the matrix. Air filled

porosity decreases as microorganisms degrade the solid matrix (Richard et al., 2000).

Substrate sterility is often needed to eliminate the growth of competing microbial

contaminants (Pandey et al., 2000). However, when the cultivated microorganism is fast growing, or produces antimicrobial compounds, substrate pasteurization might be enough to reduce the growth of contaminants, as in the case of commercial Pleurotus osteratus cultivation. Claims have been made regarding the use of hydrogen peroxide in eliminating contaminants in mushroom spawn and substrate preparation (Wayne, 2005).

However, in most cases thermal inactivation of contaminants is used.

2.7.5. Fungal Biomass Estimation in SSF

Direct estimation of fungal biomass in solid-state fermentation (complete recovery of biomass) is not possible (Raimbault, 1998). However, some studies have suggested the use of membrane culture (Mitchell et al., 1991; Nagel et al., 2000) or the use of scanning electron microscopy (Raimbault, 1998). In the case of membrane culture,

28 total biomass is removed as the membrane is peeled off the underlying substrate. In

reality this cannot be applied to commercial SSF. It is yet to be determined if microscopy

can be used as a method for direct biomass estimation.

Currently, the most widely used methods for fungal biomass estimation are

indirect methods: respirometric measurement, quantification of extracellular enzymes,

biomass component estimation (glucoasmine, nucleic acids, protein content, and

ergosterol).

Aerobic microbial activity is associated with oxygen uptake and carbon dioxide

evolution. Hence, the oxygen uptake and carbon dioxide are growth associated (not in the

case of endogenous respiration) and can be used for the estimation of biomass

(Raimbault, 1998). The amount of dry matter lost can be correlated to the amount of CO2

produced.

Extracellular enzyme quantification is another method for indirect biomass

quantification. In the case of Agaricus bisporus, the mycelial mass was directly proportional to the extracellular laccase activity (Wood, 1979). Good correlation between growth and various enzymes (cellulases, pectinases, amylases) have also produced good correlations with growth (Raimbault, 1998).

Biomass component estimation is also a method used to estimate growth. Protein, nucleic acids, glucosamine, and ergosterol are all used to estimate biomass. Fungal cells, similar to plant cells, are surrounded with a cell wall. However, the cell wall is not composed of cellulose, as in plants, but rather of chitin (Madigan et al., 2003). Chitin is a polymer of N-acetyl glucosamine. However, glucosamine level in fungal mycelium varies with age of the culture. Hence, it may not be an accurate measurement for biomass

29 growth (Raimbault, 1998). Ergosterol is the predominant fungal sterol. Both ergosterol and glucoasmine have been used to estimate the growth of A. bisporus (Matcham et al.,

1985). However, some studies have shown that the level of ergosterol depends on culture

conditions and is not always reliable (Nout et al., 1987).

2.8. Aseptic Processing of Particulate Materials

This research entails the development of an aseptic processing system for grain

and oilseed sterilization. After a brief description of starch gelatinization, the focus is

applied to the history, types, and principles of aseptic processing.

2.8.1. Gelatinization of Starch

Starch is an insoluble carbohydrate found in large amounts in cereal grains. It is composed of amylose and amylopectin, which are formed of glucose monomers. Heating grains in the presence of excess water is associated with starch gelatinization (Gomi et

al., 1998). Gelatinization is defined as the loss of order and crystalinity of starch

molecules and is accompanied by a swelling of the grain (Hosney, 1998). Water diffuses

into the grain and induces swelling of the starch granules. The moisture content within

grains is governed by the degree of starch gelatinization (Gomi et al., 1998). When rice is

kept in excess water at room temperature, the moisture content increases until 30% (wet

basis). However, when temperatures exceed 60 °C, the moisture content increases beyond

30% (Gomi et al., 1998). Generally, starch granules when placed in cool water (less than

50°C) absorb 30% of their weight in water (Hosney, 1998). Amylose starts to solubilize

at 70°C and with continued heating the amylopectin molecules also solubilize (Hosney,

1998). Wheat starch is completely solubilized at 130 °C (Hosney, 1998). Cooling

solubilized starch-water mixture results in a sticky gel (Hosney, 1998).

30 2.8.2. Thermal Processing and its Application to Food

The goal of thermal processing of food (defined as the application of heat as a means to inactivate foodborne pathogens, spoilage microorganisms and enzymes) is to ensure microbial safety and prolongation of food product shelf –life (Sarvacos and

Kostaropoulos, 2002). Various degrees of thermal processing are used in the food industry; pasteurization and sterilization are among the most important. Pasteurization and sterilization differ in their time-temperature combinations and their respective processing goals. For pasteurization, the goal is thermally to inactivate vegetative pathogenic, spoilage microorganisms, and enzymes. Sterilization takes pasteurization one step further by additionally inactivating spores (Maroulis and Saravacos, 2003). Taking milk as an example, pasteurization of milk is achieved by subjecting the milk to temperatures ranging from 60 ºC to 90 ºC for periods of 30 min to 4 sec, respectively; whereas sterilization of milk is usually done at temperatures ranging between 109 ºC to

150 ºC for periods of 40 min to 6 sec, respectively (Maroulis and Saravacos, 2003).

2.8.3. Steam-based Thermal Sterilization Processes

Steam-based thermal sterilization processes include in-container sterilization, indirect steam sterilization, and direct steam sterilization. In-container sterilization is the sterilization of solid, semisolid, and liquid foods after they have been sealed in containers made of metal or thermally stable plastic (Maroulis and Saravacos, 2003). It is traditionally used for canning and can be achieved either in batch (retorts) or in continuous sterilizers (hydrostatic sterilizers). The quality of the product from such sterilization processes is often poor because the ingredients are subjected to elevated temperature for extended periods of time (Sandeep and Puri, 2001) in order to ensure

31 sterilization at the center of the container. Indirect and direct steam heating consists of

sterilizing products (fluid food) to obtain commercial sterility by heating the flowing

products and keeping them in a holding tube at a given temperature for a period of time

until commercial sterility is achieved (Maroulis and Saravacos, 2003). Indirect steam

heating consists of using heat exchangers that transfer the heat from the steam to the

product without the steam actually coming in contact with the product. Direct steam

injection can be accomplished by injecting steam into the product or by passing a thin

layer of product in a steam chamber, also known as steam infusion (Sandeep and Puri,

2001).

2.8.4. Aseptic Processing History and Description

Pre-sterilization or sterilization before packaging of food products and subsequent

aseptic filling is not a new concept (Reuter, 1989). The first reported aseptic

process/packaging procedure was patented in the US in 1917 by Dunkley who steam-

sterilized cans and lids prior to filling with a pre-sterilized product. In 1921, Orla Jensen

of Denmark patented a process for packaging ultra-sterile milk which was termed

“Aseptic Conservation Process” (Reuter, 1989). In the USA during the 1920's, a process

called HCF (Heat-Cool-Fill) was developed in which cans and lids were sterilized in a

steam/air pressurized chamber filled with a sterile product and sealed (Reuter, 1989;

Sandeep and Puri, 2001). In 1942, a process termed the “Avoset” process was developed.

The product was sterilized with steam injection while the packages/cans were sterilized using a retort or hot air (Sandeep and Puri, 2001). During the 1940s, the Dole-Martin process was initiated and then scaled-up in the 1950s whereby Dole built the first aseptic processing/filling plant (Reuter, 1989; Sandeep and Puri, 2001). At the end of the 1940's,

32 a Swiss dairy enterprise and machinery manufacturer developed a Ultra High

Temperature (UHT) system for milk and produced the Tetra Pack System (Reuter, 1989).

In the 1960's and 1970's, the Tetra Brick and Combiblock aseptic systems were developed, respectively (Sandeep and Puri, 2001). More recently, aseptic processing was propelled by FDA approval of the use of hydrogen peroxide to sterilize packages in

1981 and the FDA’s “No Objection” letter to Tetra-Pack for the aseptic processing of low-acid food (non-alcoholic, pH > 4.6, water activity > 0.85) with large particles

(Sandeep and Puri, 2001).

Sandeep and Puri (2001) defined aseptic processing as a means to sterilize

products by heating, holding for a period of time, cooling, and finally packaging in a

sterile container. Therefore, the different steps in an aseptic processing system are Heat-

Hold-Cool-Fill.

There are two types of aseptic processing: batch aseptic processing whereby the

product is sterilized in large kettles, cooled, and then placed in sterile containers; and

continuous aseptic processing whereby pipes and heat exchangers are used to heat, hold,

cool and finally fill the product into sterile containers. The advantage of aseptic

processing and continuous aseptic processing in particular is that the product is of higher

quality because it is not subjected to prolonged periods of elevated temperatures. Other

benefits of aseptic processing are that less energy is consumed, automation is possible,

and any size/type of package can be used (Sandeep and Puri, 2001).

33 2.8.5. Elements of a Continuous Aseptic Processing System

2.8.5.1. Heating Section

Heating of the product in a continuous aseptic processing system can be achieved

by either direct steam injection/steam infusion or via non-contact heat exchangers. There are different types of non-contact heat exchangers: plate, tubular, shell and tube, and scraped surface. For heat exchangers, they can be further classified as concurrent and countercurrent heat exchangers (Sandeep and Puri, 2001).

2.8.5.2. Holding Section

For aseptic processing of foods, the FDA has mandated that the residence time in the heating section cannot contribute to lethality. Only the residence time within the holding section can contribute to lethality (Sandeep and Puri, 2001). In addition,

residence time within the holding section should be determined using the mass flow and

specific volume of the product and not the pump displacement (Sandeep and Puri, 2001).

2.8.5.3. Cooling Section

Once the product is sterilized in the holding section, it is cooled down rapidly to

an appropriate temperature. Juices, milk, and other low viscosity food products are cooled down to 20 °C; whereas high viscosity products such as puddings are cooled to

40ºC (Reuter, 1989). Zhang and Sun (2005) evaluated different methods for cooling

cooked rice in which they concluded that vacuum cooling (compared to blast cooling,

plate cooling, and cold room cooling) was the most effective method of cooling cooked

rice from an average temperature of 80-90 °C down to 4 °C in 4 min. Final moisture

content of the vacuum cooled rice was significantly lower than other methods of cooling

(reduction of 11.4%).

34 2.8.5.4. Aseptic Filling and Packaging Section

To ensure that the sterile product is not contaminated in the filling machine, the container surfaces and the processing environment have to be sterile (Reuter, 1989).

Various methods such a thermal methods, irradiation, and chemical methods are used to sterilize surfaces. Thermal methods consist of steam, hot air, and mixtures of hot air and steam. Irradiation methods consist of irradiating with infrared, ultraviolet, or ionizing rays. Chemical methods include washing surfaces with hydrogen peroxide, peracetic acid, and others.

2.8.6. Liquid-based Segmented Flow Aseptic Processing System

The segmented-flow system, conceptually characterized by Stephens and Walker

(2003) as a continuous “series of cans end-to-end going through a heat-hold and cool cycle”, can handle food with various particle sizes. The main advantage of the segmented flow system is the ability to control liquid and particle residence time simultaneously.

The traditional pipe-flow system is replaced by a pipe containing a chain connected to disk barriers which compartmentalize set volumes of liquid/solid medium (Stephens and

Walker, 2003). The chains are connected to a sprochet which in turn is connected to a drive motor. Varying the speed of the motor translates to varying the volumetric flow rate of the product which affects the residence time of the fluid/particle medium (Walker and

Beelman, 2002). Figure 2.1 shows a diagram of the segmented flow aseptic processing system.

35 Drive Motor

~20 psi Steam or Sterile

Compressed Gas

Refrigeration

Backpressure Pump (level controlled)

Product

Boiler

Feed Feed Pump (level controlled)

Figure 2.1. Segmented-flow aseptic processing schematic (Walker and Beelman, 2002).

2.8.7. Steam-based Segmented-flow Processing System

The segmented-flow technology was again used in the continuous steam aseptic processing system, specifically for particulate matter that is subject to mechanical damage (Anderson and Walker, 2005). A U-cross-section conveyor-bed system was designed and constructed in an 20.3 cm sanitary tube to transport the particulate matter.

The variable drive on the conveyor-belt system enables control of the residence time of the particles within the sterilization chamber for heating and holding times. Rather than having liquid in the sterilization section, steam is directly applied to the 15.3 cm sanitary tube which totally surrounds the particles on the conveyor belt system. A temperature of

131°C inside the sterilization chamber was achieved. Figure 2.2 provides a diagram of the continuous-steam, segmented-flow, aseptic processing system.

36

Figure 2.2. Continuous steam segmented-flow, aseptic processing system (Anderson and Walker, 2005)

2.9. Continuous Production of Mushroom Spawn

Holtz and McCulloch (1995) developed and patented a process in which grain spawn was produced continuously. The system was composed of an aseptic processing system and several 100 L airlift bioreactors. The aseptic processing system, a hollow auger type screw, was used to sterilize the grain. An additional section composed of an auger cooling screw was used to reduce grain temperature by vacuum cooling. The sterilized grain was inoculated with a sterile broth of mushroom mycelium produced in the bioreactors. The grain and mycelium were mixed thoroughly and aseptically filled in sterile plastic bags.

37 2.10. Summary of Literature Review

Agaricus bisporus is primarily grown on a composted substrate that is

increasingly causing problems for mushroom producers because of the environmental

impact large-scale composting activities are causing on neighboring residents of

mushroom farms. There are two main areas of research that have/are attempting to

address this problem. The first focuses on developing methods to reduce the impact of

composting activities on the environment, whereas the second focuses on eliminating

compost from the substrate. Developing and refining non-composted substrates is not a

new idea, as there is extensive research on substrates composed of various agriculture waste products. Grain-based substrates were shown to be a suitable substrate for A.

bisporus mushroom production but little work has focused on developing and refining

such substrates for increase mushroom production and substrate bioefficiency.

Additionally, the literature yielded very little work on designs for alternative mushroom production systems adapted for non-composted substrates.

Grain processing is a mainstay in the mushroom spawn industry. However, the current systems employ batch sterilization methods, whereby all the ingredients are added

to large vessels, sterilized, and then inoculated with the mushroom fungus. Adapting

continuous steam sterilization processes such as aseptic processing units based on

segmented-flow technology will enable continuous production of grain-based substrates a

clear advantage over batch sterilization processes.

38 Chapter 3

Production of Agaricus bisporus Mushrooms on Commercial Grain Spawn Mixed with S41 and S44 Supplements

3.1 Abstract

Non-composted substrates composed of commercial grain spawn and delayed- release nutrient supplements were tested for mushroom production as alternatives to the

environmentally problematic composting process associated with conventional

commercial mushroom (Agaricus bisporus) cultivation. The effect of casing type,

thiophanate-methyl (fungicide) use, delayed-release supplement type (S41 and S44) and

rate, and perlite addition were tested on mushroom production. Use of a non-sterile

casing overlain on grain spawn/supplement substrate produced mushrooms comparable to a sterile casing with 25% activated carbon, yielding 6.4 kg/m2 and 7.6 kg/m2,

respectively. Thiophanate-methyl was used in one set of treatments to control the incidence of fungal contamination. However, mushroom yield was severely reduced.

Average mushroom yield from treatments with S41 supplement was greater than that of

S44. When the commercial grain spawn substrate was underlain with a layer of perlite

(2000 ml over 0.048 m2) to absorb and release excess water, mushroom yield increased

dramatically, producing 13 kg/m2 compared to 7.6 kg/m2 for treatments without perlite.

Keywords: button mushroom, activated carbon, non-composted substrates, perlite

39 3.2. Introduction

Cultivation of Agaricus bisporus, the button mushroom, is widely practiced

around the world, with annual sales in the U.S. generating more than $841 million (Rai,

2004; USDA, 2005-2006). Traditional mushroom production, a solid substrate

fermentation process (Miller, 1996), involves a series of steps (composting, spawning,

casing, pinning, and harvesting) universally adopted by mushroom producers with only

slight variations.

Composting within the mushroom production process is typically divided into two

steps, Phase I and Phase II (i.e. short method), generating a partially composted substrate

colonized by thermophilic microorganisms conferring selectivity for the growth of A. bisporus (Schisler, 1982; Straatsma et al., 1994). This substrate preparation method has

come under increasing scrutiny mainly due to the production of unpleasant odors and

nutrient rich runoff from mushroom substrate preparation sites (Derikx et al., 1990;

Heinemann and Wahanik, 1998). Furthermore, after being used in mushroom production,

the spent mushroom substrate (SMS) is discarded in large volumes and placed in fields

where it resumes composting and releases salt-rich leachate that may enter groundwater

or surface water runoff (Guo et al., 2001; Heinemann et al., 2003). Beyer (2006)

estimated that two of the main mushroom production counties in Pennsylvania produced

580,000 m3 of SMS annually. Aside from the environmental aspect, the composting

process is time consuming, taking up to three weeks for preparation, and both labor and

machinery intensive, necessitating periodic watering, turning, and handling.

Although not always clearly defined, there are two general research approaches

addressing the environmental issues created by the composting stage of mushroom

40 production. One approach focuses on developing different strategies and designs to reduce odor production and nutrient runoff from composted substrate and SMS, such as aerating composted windrows, using microporous membranes, and indoor composting

(Labance et al., 1999; Rai, 2004). The other is a more radical approach , investigating the use of non-composted substrates for mushroom production. The latter approach would result in the elimination of the composting stage from mushroom production (Till, 1962;

San Antonio, 1971; Mee, 1978; Sanchez and Royse, 2001; Bechara et al., 2006ab).

Mamiro et al., (2007) showed that a combination of spent mushroom compost mixed with a non-composted substrate, based on a formulation used by Sanchez and Royse (2001), was also a viable alternative that would reduce, but not eliminate, the composting from mushroom substrate preparation. A better understanding of the current state of the art of traditional compost-based mushroom production systems will help improve mushroom yield in non-composted substrates.

Mushroom yield on composted substrates has increased tremendously because of the development of hybrid varieties of A. bisporus, improvements in the composting method, and the use of delayed-release supplements (Sonnenberg, 2000). To increase mushroom yield in composted substrates, delayed-release nutrient supplements consisting of oilseed or proteinaceous compounds are added to the substrate, either at spawning or at casing (Carroll and Schisler, 1976; Nair et al., 1993; Romaine and Marlowe, 1993).

Published data showed that 2-20% increases in mushroom yield are possible with the addition of delayed-release nutrient supplements (Schisler and Patton, 1972; Randle,

1983).

41 Growth of the mushroom fungus on cereal grains is a widely used process for the production of mushroom grain spawn, the vehicle used to inoculate substrates, which was

initially developed by Sinden (1932). In some instances, the grain spawn is coated with a

mixture of calcium carbonate and thiophanate-methyl used for the control of the fungal

pathogen Trichoderma aggressivum f. aggressivum just before it is added to the substrate

(Royse and Romaine, 2002). Kananen et al. (2000) developed and patented a grain spawn

substrate composed of grains and supplement mixtures (both delayed-release and protein

rich supplements) that was termed “mushroom spawn-supplement”. The average yield of

mushrooms from a compost-based substrate inoculated with the spawn-supplement

mixture was greater than a similar substrate inoculated with traditional rye grain spawn.

Apart from greater mushroom yield, Kananen et al. (2000) also reported lower incidence

of disease and shorter spawn runs (rapid colonization of the composted substrate) for

substrates inoculated with mushroom spawn-supplement.

Casing, which is the addition of a layer of neutralized peat moss over the

mushroom substrate, is an essential step for the fruiting of the A. bisporus. The exact role

of the casing layer in mushroom development has yet to be determined. However, several

hypotheses have been proposed, such as creating a gradient of carbon dioxide or of an

unidentifiable volatile, supplying the mushroom fungus with essential microorganisms

that induce fruiting, and/or removing a specific mushroom inhibitor (Tschierpe, 1959;

Eger, 1961; Peerally, 1978; Noble et al., 2003). One consistent observation has been that

sterilized casing produced few mushrooms and, in some cases, mushroom fruiting was

completely inhibited. However, adding activated carbon (AC) as part of the casing

42 material or as the casing material itself improved or restored mushroom yield (Long and

Jacobs, 1974; Verbeke and Overstyns, 1991; Noble et al., 2003).

The widespread success of growing the vegetative stage of the mushroom fungus

on cereal grains might have compelled San Antonio (1971) to develop a non-composted

substrate for A. bisporus mushroom production using rye grain as substrate. The substrate

formulation used in that study was as follows: 200 g of rye, 20 g of CaCO3, and 200 ml of de-ionized water. Although the study did not present yield data, the author concluded that mushroom yield was comparable to compost-based substrates. More recently,

Bechara et al. (2006ab) developed different non-composted grain based substrates for A. bisporus mushroom production. One substrate tested was commercial mushroom grain

spawn with different proportions of perlite, a granular siliceous material that can hold

four times its weight in water. The highest mushroom yield obtained for the commercial

grain spawn substrate was 5.3 kg/m2, whereas yield from composted substrates was 7.7

kg/m2. The authors concluded that the vegetative growth of the mushroom fungus

exhausted the nutrient reserves within the grain. One area not previously explored is the

use of grain-based substrates supplemented with delayed-release nutrient supplements,

typically applied to partially composted substrates.

The goal of the current study was to test several factors that influence mushroom

yield in a non-composted substrate composed of a mixture of commercial grain spawn

and delayed-release nutrient supplements. The first factor involved the type of casing:

sterile and non-sterile. The second set of factors tested the effect of coating the grain

spawn with thiophanate-methyl and supplementing the substrate with two different

soybean-based delayed-release nutrient supplements. Finally, the addition of a water-

43 holding material, horticultural-grade perlite, was assessed for its effect on mushroom

production.

3.3. Methods

3.3.1. Mushroom Grain Spawn, Nutrient Supplement and Water-holding Material

Commercial off-white hybrid mushroom rye grain spawn A15 (Sylvan Spawn

Laboratories, Kittanning, PA) was used in this study. Two delayed-release nutrient supplements, S41 and S44 (Full House- Beta Spawn Co. Inc., Toughkenamon, PA), were also used. Both supplements were composed of cracked soybean coated with different proprietary antimicrobial films. In some treatments, horticultural-grade perlite (Therm-o-

Rock East, New Eagle, PA), a water-holding material that can hold four times its weight

in water, was used.

3.3.2. Preparation of Mushroom Production Containers

Sterilized (121°C for 60 min) polypropylene containers (L = 0.3 m, W = 0.16 m,

D = 0.09 m) were used as mushroom production containers. When perlite was used, it

was saturated with tap water and then added to the containers prior to autoclaving. The

ingredients of the mushroom substrate were commercial rye grain spawn and S41 or S44

nutrient supplement. The total biological matter, spawn + S41 or S44, was standardized

to 800 g/container (wet basis). When thiophanate-methyl (Topsin®, Cerexagri Inc., King

of Prussia, PA) was used, it was coated onto the grain spawn with calcium carbonate as a

bulking agent using the following ratio: 0.188 g Topsin/150 g spawn/9.3 g CaCO3. A

0.015 m casing layer (limed peat moss), as normally prepared by the Penn State

University (PSU) Mushroom Test Demonstration Facility, was overlain on the spawn and

S41 or S44 mixture. Sterilized casing was prepared by mixing the casing with 25%

44 volume/volume (v/v) AC (Fisher Scientific, Hampton, NH) and autoclaving the mixture

(121°C, 120 min). The pH of the casing increased from 6.5 to 7.5 with the addition of the

AC, with no further change in pH measured post-autoclaving.

3.3.3. Mushroom Production Chamber

The filled containers were transported to a mushroom production chamber at the

Mushroom Research Center (Penn State). All trials, irrespective of treatments, used

chamber set points of 22 ºC and 80% RH. The casing layer was kept moist by daily

watering with regular tap water using a rose-face nozzle. Once the mycelium covered

50%-70% of the casing surface in all of mushroom production containers (typically 10

days after placing in chamber), pinning was induced by reducing the temperature to 16 ºC

and increasing chamber ventilation to achieve a carbon dioxide concentration of 800

ppm. Mushroom harvest began typically 21 days after entering the production chamber.

The mushrooms were harvested manually for a three-week period, and weight was

recorded immediately after harvest.

3.3.4. Summary of the Experimental set-up

Different factors were evaluated for their effect on mushroom production. The

first experiment evaluated two different casing treatments: a non-sterile casing and a

sterile casing containing 25% AC (v/v), each placed over a grain substrate composed of

commercial rye grain spawn and 10% S41. The second set of experiments compared

mushroom yield from substrates composed of commercial grain spawn and S41 or S44

supplements added at different levels (0%, 1%, 5%, 10%, 20%), and the effect of coating the grain spawn with thiophanate-methyl. The third and final experiment evaluated the

45 influence of adding a 2000 ml (0.042 m) layer of perlite beneath a commercial grain spawn substrate containing 10% S41.

3.3.5. Parameters Evaluated and Analysis of Data

All treatments were replicated three times. The results were presented as yield/unit area (kg/m2), and bioefficiency as a percentage (fresh weight of mushroom/dry weight of substrates).

Yield and bioefficiency data were statistically analyzed using General Linear Model for multiple comparisons, whereas a two-sample t-test was used to compare experiments composed of two treatments. For comparisons among treatments within an experiment, the Tukey method for pairwise comparisons (Tukey, 1949) was used to determine statistical differences among treatment means. MINITAB Statistical Software Package

(Release 13.1, State College PA) was used for the statistical analysis.

3.4. Results

The effect of casing treatment on A. bisporus mushroom yield was not significant

(p>0.05). On average, the yield from casing with 25% AC was 7.6 kg/m2, whereas yield from non-sterile casing was 6.4 kg/m2. Table 3.1 summarizes the data for the casing treatment experiment. A sterilized casing with AC was adopted for all other treatments to ensure that microbial contaminants did not affect yield.

Table 3.1. Mushroom yield from two different casing treatments, with and without AC Mushroom Yield Bioefficiency Treatment (kg/m2)1,2 (%) Sterile casing + 25% AC 7.6 ± 1.6a 78.4 ± 16.2 a Non-sterile Casing 6.4 ± 3.6a 66.6 ± 37.2 a 1 Data presented with mean ± standard deviation 2 Treatments followed by similar letters are not significantly different (p<0.05 is significant)

46 Mushroom yield from treatments with grain spawn coated with thiophanate-

methyl, regardless of nutrient supplement type (S41 or S44) used, was lower than those

of treatments with untreated grain spawn (p<0.05). Overall, there was no significant

effect of supplement type on mushroom yield for treatments with thiophanate-methyl

(p>0.05). However, rate of supplementation was significant (p<0.05), with an observed

decrease in yield and bioefficiency for treatments with higher rates of S41 and S44.

Complete inhibition of mushroom formation occurred at 20% S44, and similarly, a yield

of only 0.46 kg/m2 was produced in the 20% S41 treatment. The highest recorded yield

for treatments coated with thiophanate-methyl was produced by the 5% S41 treatment

(2.51 kg/m2) and this was the only treatment that was significantly different from all other treatment means. Results of treatments with thiophanate-methyl are presented in

Table 3.2.

Table 3.2. Mushroom yield from grain spawn substrates treated with thiophanate-methyl and supplemented with S41 and S44. Mushroom Yield Bioefficiency Treatment (kg/m2)1,2 (%) 0% S41and S44 1.51 ± 1.00 a,b 16.8 ± 11.9 a,b 1% S41 1.98 ± 0.90 a,b 21.9 ± 10.7 a,b 5% S41 2.51 ± 0.50 b 26.9 ± 5.4 b 10% S41 0.83 ± 0.13 a,b 8.7 ± 1.3 a,b 20% S41 0.46 ± 0.25 a 4.5 ± 2.4 a,b 1% S44 1.45 ± 0.87 a,b 15.9 ± 9.6 a,b 5% S44 1.50 ± 0.41 a,b 16.4 ± 1.5 a,b 10% S44 0.81 ± 0.77 a,b 8.5 ± 8.0 a,b 20% S44 0.00 ± 0.00 a 0 ± 0 a 1 Data presented with mean ± standard deviation 2 Treatments followed by similar letters are not significantly different (p<0.05, is significant)

For treatments without thiophanate-methyl, supplement type and rate of

application had a significant effect on mushroom yield and bioefficiency (p<0.05).

Although not statistically significant, the highest yield of mushrooms from treatments

47 without fungicide application was produced on the grain spawn substrate supplemented

with 5% S41 (9.66 kg/m2). Although not statistically significant, the highest mushroom

yield among the S44 treatments was the grain spawn substrate supplemented with 20%

S44 producing 4.03 kg/m2. Average yield from S41 treatments was higher than yield from the S44 treatment. The same conclusion can be drawn for the bioefficiency values.

Table 3.3 summarizes the results for treatments without thiophanate-methyl coated spawn.

Table 3.3. Mushroom yield from grain spawn substrates without thiophanate-methyl application supplemented with S41 and S44 Mushroom Yield Bioefficiency Treatment (kg/m2)1,2 (%) 0% S41and S44 1.38 ± 0.97 a 15.3 ± 10.9 a 1% S41 5.42 ± 1.46 b,c,d 59.7 ± 16.2 b,c,d 5% S41 9.66 ± 0.97 e 103.4 ±10.4 e 10% S41 7.56 ± 1.56 c,d,e 77.9 ± 16.1 c,d,e 20% S41 8.63 ± 2.26 d,e 83.0 ± 21.8 d,e 1% S44 2.91 ± 1.62 a,b 32.0 ± 17.9 a,b 5% S44 1.74 ± 1.55 a,b 18.6 ± 16.5 a,b 10% S44 1.98 ± 0.28 a,b 20.4 ± 2.9 a,b 20% S44 4.03 ± 0.96 a,b,c 38.7 ± 9.2 a,b,c 1 Data presented with mean ± standard deviation 2 Treatments followed by similar letters are not significantly different (p<0.05, is significant)

The inclusion of a 4 cm underlayment of perlite markedly increased mushroom

yield from 7.56 kg/m2 to 13.01 kg/m2, with biological efficiencies of 76.1% and 131%

respectively (Figure 3.1).

48 16.00

14.00

12.00

10.00

8.00

Yield (kg/m2) 6.00

4.00

2.00

0.00 No Perlite-10% S41 Perlite-10% S41

Figure 3.1. Effect of adding a single layer of perlite below the commercial grain spawn substrate containing 10% S41 on mushroom production

3.5. Discussion

The composting process within the mushroom production framework creates a complex “living” substrate that is inherently specific for the growth of A. bisporus.

Microbial contaminants that are unwillingly introduced into the composted substrate due

to the addition of non-sterile casing or through other means are kept in check by

competitive exclusion. When using sterilized substrates, such as grains, this specificity is

lost, allowing competing microorganisms to overrun the growth of A. bisporus (Ross and

Harris, 1983). Commercial grain spawn is typically rye or millet entirely colonized with a

“barrier” of mushroom fungus mycelium that protects the nutrient-rich reserve of the

grains. This “barrier” can be likened to the specificity of composted substrates to A. bisporus growth. However, when adding delayed-release nutrient supplements to grain spawn, and in doing so forming grain spawn-supplement substrates, there is substantial risk that un-colonized delayed-release nutrient supplements become a readily available nutrient source for growth and spread of contaminants within such substrates. This is why

49 it was important in this research to compare the effect of non-sterile casings to sterilized

casing in such non-composted substrates, recognizing that previously cited research demonstrates that the addition AC to sterilized casing is a must. From yield data, there was no statistical difference between sterilized casing with 25% AC and non-sterile casings lacking AC. However, the variation in yield for the non-sterile casing was greater than that for the sterile casing with AC. This finding concurs with the results of an earlier study on composted substrates cased with non-sterile casing and sterile casing with AC

(Bechara et al., 2006ab). This increase in variation among replicates of a comparable substrate with non-sterile casing is believed to be due to the growth of opportunistic microbial contaminants. The findings of a recent study by Chikthimmah et al. (2006) indicated that both sterilized and pasteurized casing may lead to unimpeded growth of human pathogens, such as Salmonella sp., which in non-sterile casings are inhibited by the naturally occurring microflora. This raises the question whether casing should be heat-treated at all. However, for research purposes, the use of sterilized casing can be used as a tool to reduce variation among replicates, which is the basis for its use in the present study.

Trichoderma sp. and other fungi are serious competitive contaminants that can

severely reduce mushroom yield in commercial production. Treating the grain spawn

with a fungicide, such as thiophanate-methyl, was thought to reduce the incidence of

fungal contamination and increase yield. Treatments involving the addition of

thiophanate-methyl to the grain spawn using the same dose of fungicide did not support

this hypothesis. The results clearly show that the fungicide, along with CaCO3 as a

bulking agent, at the given dosage severely reduced mushroom yield in non-composted

50 commercial grain spawn substrates. This inhibitory effect, not apparent in commercial

settings, is probably attributed to the bulking effect of compost where only 1.5% of grain

spawn/dry weight of compost is used for inoculation. When the substrate is almost entirely grain spawn, lower fungicide dosages should be tested to determine optimal levels that would protect the substrate from fungal contaminants.

The addition of delayed-release nutrient supplements S41 and S44 had an effect on mushroom yield. The S44 supplement in all treatments yielded less mushrooms compared to the S41. Furthermore, mold growth was detected in all treatments

supplemented with S44 nutrient supplement, but not so with the S41. Both are soybean-

based supplements differing in a few proprietary ingredients added to the coating.

Mushroom yield from the 20, 10, and 5% S41 treatments were comparable (p>0.05).

Since the unit price of S41 is higher than for grain spawn, 5% S41 spawn substrate would

be the most economically viable substrate. For the S44 mushroom nutrient supplement,

differences in mushroom yield for the 10%, 5%, 1% S44 and 0% were not significant

(p>0.05). This was due to the large variation across replicates for each treatment.

The impetus for testing the addition of perlite, a water-holding material, on

mushroom yield was two-fold. First, it was explored to determine if adding a water

reservoir to maintain, and perhaps increase, the moisture content of the substrate would

translate to higher yield. Generally, when partially composted substrates are used, the

final moisture content at spawning is around 68%. The grain spawn contains 40-48%

moisture (wb) depending on supplier, type of grain, and age, so it seems that adding a moisture reservoir might be beneficial. Second, the perlite layer absorbs excess water

from the casing that percolates through the substrate and obviates the otherwise water-

51 logging and rotting of the cereal grain substrate. Adding the perlite raises the grain bed

depth and absorbs the extra water that would have collected at the bottom. It is believed

that the perlite acts as both a source and sink for water. This hypothesis (source and sink)

is supported by the tremendous increase in mushroom yield obtained from substrates

placed over a single stratum of perlite. These substrates produced 13 kg/m2 average yield

with a bioefficiency of 131% compared to 7.6 kg/m2 with a bioefficiency of 76% for

similar substrates with no perlite. However, yield was still lower than mushrooms

produced on partially composted substrate (28.9 kg/m2). Typically, composted substrate

depths in the range of 0.16-0.20 m with measured bioefficiency from 80 to 90% are

considered good (Schisler, 1982), whereas depth of the commercial grain spawn/supplement substrate used here ranged from 0.02-0.025 m with a maximum recorded bioefficiency of 131%. It is anticipated that increasing the non-composted

substrate bed depth would directly translate to an increase in yield.

3.6. Conclusions

A non-composted substrate composed of a mixture of commercial grain spawn

and a delayed-release nutrient supplement successfully produced A. bisporus mushrooms.

Mushroom yield from a sterile casing with 25% AC yielded comparably to a non-sterile casing alone with less variation among replicates. Hence, the former was used for all the subsequent treatments. The addition of the fungicide thiophanate-methyl to the grain substrate severely reduced mushroom yield. The addition of delayed-release nutrient

supplement S41 yielded more mushrooms than S44 for both treatments with and without thiophanate-methyl. Finally, including a layer of perlite beneath the grain-based substrate increased mushroom yield from 7.5 kg/m2 to 13 kg/m2; a significant improvement but

52 still comparatively lower than typical yields from commercial values of 28-30 kg/m2.

Finally, studies testing a wider variety of delayed-release nutrient supplements and water- holding materials are needed to further improve mushroom yield.

Acknowledgments

The authors would like to acknowledge the assistance of manager Tom Rhodes and the entire crew at the Penn State Mushroom Research Center. Funding for this study was provided by the College of Agricultural Sciences and the Department of Agricultural and Biological Engineering (Penn State). The grain spawn was generously supplied by

Mark Spear from Sylvan Spawn Labs.

53 Chapter 4

Factors Influencing Mushroom Yield in Non-composted Commercial Grain Spawn Substrates

4.1. Abstract

Mushroom substrates composed in part of commercial grain spawn supplemented with S41 (delayed-release nutrient supplement), overlaid on a bed of water-holding materials, were evaluated for mushroom yield (Agaricus bisporus) and substrate

bioefficiency (BE). Water-holding material type (i.e. perlite, vermiculite, sand,

polyacrylamide) and volume, S41 rate, casing material, and grain spawn types were

varied. There were no significant differences for yield and BE among treatments in which

the type of water-holding material was varied (p>0.05). However, average mushroom

yield was highest for the perlite treatment, which produced 8.68 kg/m2 with a BE of

102.9%. Increasing S41 beyond a 5% rate did not significantly increase mushroom yield

(p>0.05). Mushroom production was greatest for a sterilized casing layer with activated

carbon (AC) compared to that of treatments receiving a non-sterilized casing layer

(p<0.05). The effect of varying the volume of perlite was significant for treatments

receiving S41with the highest yield corresponding to the 1000-2000-ml application

(p<0.05). BE peaked for treatments receiving a 2000 ml perlite amendment. The highest

recorded yield of 14.28 kg/m2 and a BE of 177% was observed in the 5% S41 + 2000 ml

perlite + sterilized casing with AC treatment. Finally, there was no significant difference in mushroom yield for the two types of commercial grain spawn (millet and rye) used as the basal mushroom substrate with an S41 amendment and a perlite underlay (p>0.05).

These findings suggest that adding a layer of perlite and using sterilized casing with AC

54 will increase mushroom yield compared to treatments with unsterilized casing and no

perlite for a non-composted substrate composed of commercial grain spawn and a

delayed-release nutrient supplement (S41).

Keywords: Agaricus bisporus, activated carbon, solid state fermentation

4.2. Introduction

Commercial production of Agaricus bisporus, the button mushroom, is achieved

using a system that relies on composting of plant and animal matter (horse-bedded manure, hay, corn cobs, among others) to produce a matrix that is both selective to A. bisporus growth, and supports high yielding mushroom crops. Mushroom yield from commercial (composted) substrates may vary depending on region. However, the national average for the U.S.A. is 30 kg/m2 (USDA, 2005-2006). Composting within the

mushroom production process has come under intense scrutiny, mainly due to odor

production and nutrient-rich run-off /leachate during substrate preparation and disposal

phases (Derikx et al., 1990; Guo et al., 2001; Heinemann and Wahanik, 1998;

Heinemann, et al. 2003).

Composting, as a substrate preparation method, is not an intrinsic step for A.

bisporus mushroom production (San Antonio, 1971; 1975). Several studies showed that

A. bisporus is capable of producing mushrooms on non-composted substrates. One of the

earliest studies was undertaken by Till (1962) in which he proposed the use of sterilized

substrates predominantly composed of straw and other lesser ingredients. Huhnke and

Von Sengbush (1968) proposed the use of pasteurization as opposed to sterilization for

the preparation of a mushroom substrate. Although the authors report mushroom yields

being comparable to composted substrate, they also report that yields were highly

55 variable. Other studies on developing non-composted substrates have used leached cow

manure, rye grain, and substrate mixtures composed of sawdust, seeds, and other lesser

ingredients (Mee, 1978; San Antonio, 1971; Sanchez and Royse, 2001).

Grain-based non-composted substrates were first reported by San Antonio (1971)

in which sterilized rye grains colonized by A. bisporus were used as the basal substrate

for mushroom production. Typically, the seeding material used in inoculating composted

substrates is sterilized rye or millet grain that are fully colonized with the vegetative mycelium of the mushroom fungus. The product is termed “ mushroom grain spawn”.

When grain spawn is added to composted substrates, the mycelium colonizes the entire substrate. San Antonio (1971) was successful in showing that grain spawn can be used as

a substrate for mushroom production. Bechara et al. (2006a) tested the effect of using a non-composted substrate composed of commercial rye grain spawn mixed with different

rates of perlite, a water-holding material, on mushroom production. The authors state

that adding perlite to the grain spawn substrate triggered mushroom production a week

earlier than treatments lacking perlite. Bechara et al. (2005ab) also showed that adding

commercial delayed-release nutrient supplements, such as S41, typically added to

compost-based substrates to improve yield, also increased yield in non-composted grain

spawn substrates (9.7 kg/m2) compared to no supplement addition (1.4 kg/m2). Adding

perlite as a single layer below the grain spawn/delayed-release nutrient substrate increased yield from 7.5 kg/m2 to 13 kg/m2. It was concluded that the perlite layer raises

the grain substrate bed and probably absorbs excess water that collects at the bottom of

the mushroom trays. Additionally, it was thought that the perlite layer might act as a

water reservoir for the mushroom fungus. For the production of grain spawn, moisture

56 contents should not exceed 55% typically, otherwise “wet spots” can develop in which

the fungus fails to colonize the substrate (Stamets, 2000a). Since moisture content of

commercial grain spawn ranges from 43% to 50%, the addition of an underlying layer of

water-holding material, such as perlite, likely increases water availability for the

developing mushrooms, without the need for increasing moisture content of the grain

spawn substrate. This is especially important knowing that the mushroom fungus receives

54-83% of the water requirement for mushroom production from the substrate, and 17-

46% from the casing (Kalberer, 1990).

Spreading a top-dressing layer, termed mushroom casing, over both composted

and non-composted A. bisporus substrates is essential for Agaricus mushroom production

(San Antonio, 1975). The casing layer can be composed of different materials such as

spent mushroom compost, lime-neutralized peat moss, soil, among others (Schisler,

1982). Sterilizing mushroom casing reduces mushroom yield. However, this effect is mitigated by the addition of activated carbon (AC) to sterilized casing (Eger, 1972; Long and Jacobs, 1974; Nobel et al., 2003). Sterilizing casing, as applied to experimental trials, effectively eliminates unwanted microbial contaminants that can increase variability in

mushroom yield (Bechara et al., 2006a). However, most casing in commercial mushroom

production operations is not sterilized. Hence, it is important to establish any differences

that might arise between non-sterilized and sterilized casing on mushroom yield.

To date, studies have reported the effect of commercial rye grain as the basal

substrate for mushroom production, but the use of commercial millet grain spawn has not

been tested. It is anticipated that mushroom yield from commercial rye grain spawn

would differ from commercial millet grain spawn because of differences in

57 endosperm/grain size (San Antonio, 1971), whereby energy reserves in an individual rye

grain spawn are greater than millet grain, thereby sustaining the mycelium for longer

periods of time (Fritsche, 1988).

The present study addressed whether or not different types of water-holding

materials placed as a single layer underneath a grain spawn substrate had an effect on

mushroom yield and production bioefficiency. Furthermore, the effects of different S41

rates, depths of water-holding materials, spawn grain type, and casing treatments on

mushroom yield were determined.

4.3. Methods

4.3.1. Mushroom Hybrid Nutrient Supplement and Water-holding Material

Commercial off-white mushroom hybrid rye A15 juvenile commercial grain spawn (Sylvan Spawn Laboratories, Kittanning, PA) was used for most treatments except for treatments calling for millet grain spawn. Juvenile commercial grain spawn is spawn that is procured from the supplier without any intermediate refrigeration or hardening phase. The delayed-release nutrient supplement used in this study was S41 (Full House,

Bet Spawn Co Inc., Toughkenamon, PA). Four different water-holding materials were tested for their effect on mushroom yield: sand, perlite (Therm-o-Rock, New Eagle, PA), vermiculite (Thermo-o-Rock, New Eagle, PA) and polyacrylamide (JRM Chemicals,

Cleveland, OH)

4.3.2. Preparation of Mushroom Production Containers

Autoclaved (121°C for 60 min) polypropylene containers (0.3 x 0.16 x 0.09 m) were used for mushroom production. When a water-holding material was used, regardless of type, it was fully wetted with tap water, allowed to drain gravitationally and then

58 added to the containers prior to autoclaving. The containers were covered with aluminum

foil and autoclaved for 45 min (121°C and 103.4 kPa) and then allowed to cool. The

ingredients of the mushroom substrate were defined as commercial grain spawn and S41

nutrient supplement. The total biological matter, commercial grain spawn + S41, was 800 g/container (wet basis). Hence, a 0% S41 treatment corresponded to 800 g of commercial

grain spawn, whereas a 10% S41 treatment corresponded to 720 g commercial grain

spawn and 80 g S41. A casing layer (limed peat moss), as prepared by the Mushroom

Test and Demonstration Facility (The Pennsylvania State University), when sterilized,

was autoclaved (121 °C and 103.4 kPa) after adding 25% (v/v) activated carbon (AC)

(Fisher Scientific, Hampton, NH) to the material. Sterilized and non-sterilized casing

with activated carbon were spread evenly over the commercial grain spawn substrate.

Mushroom production was carried out in a chamber at the Mushroom Research Center

(The Pennsylvania State University) in which temperature and relative humidity (RH)

were controlled.

4.3.3. Environmental Conditions in Mushroom Production Rooms

The temperature in the mushroom production room was 22 °C and an atomizing

humidifier was used to supply moisture to the air (80-90% RH). Mushroom production

containers were watered on a daily basis using a rose-faced nozzle. Once 50% of the

casing surface area was colonized by mycelia (corresponding to 10 days after containers

were set in chamber), temperature was reduced to 16 °C. Mushroom harvesting for all

treatments started on average 21 days after placing containers in the production room.

Mushroom weight was recorded immediately after harvest. Harvesting was completed

three weeks beyond the first day of harvest, making for 42-day experimental period.

59

4.3.4. Experimental Set-up and Analysis of Data

The first set of treatments tested the effect of different water-holding materials

(perlite, vermiculite, sand, polyacrylamide, 1/1 (v/v) mixture of polyacrylamide and

perlite) placed as a single stratum underneath commercial grain spawn substrate

supplemented with 10% S41. The depth of the various water-holding materials was 0.042

m corresponding to a volume/container of 2000 ml for all treatments. Sterilized casing

with activated carbon was used for all treatments. Based on the results of these pilot

experiments, the water-holding material that produced the highest yield was adopted for

subsequent experiments.

The second set of treatments varied the volume of the water-holding material (0,

1000, 2000, 3000 ml) corresponding to the following depths within the mushroom

production trays (~0, 0.021, 0.042, and 0.063 m), the rate of S41 addition (0, 5, 10, 15,

and 20%), and the casing (sterilized and non-sterilized). Simultaneously, an experiment

was set up that varied the type of commercial grain spawn (millet and rye) and S41 rate

(0, 5, 10, 15, and 20%) and included the use of sterilized casing with 25% AC with an

underlying layer of 1000 ml perlite.

All treatments were replicated three times. The results of the treatments were

presented in terms of mushroom yield/unit area (kg/m2) and BE (% fresh weight of

mushroom/dry weight of substrates). For the first experiment, a one-way ANOVA was used to statistically analyze the data at α = 0.05. For the other experiments, yield and BE

data were statistically analyzed using the General Linear Model with α = 0.05. MINITAB

60 Statistical Software Package (Release 13.1, State College, PA) was used for the statistical

analysis.

4.4. Results

There were no significant differences in mushroom yield among the different

types of water-holding materials tested. Yields for perlite, vermiculite, sand, polyacrylamide, and polyacrylamide + perlite substratum (1/1) treatments were not

significantly different (p>0.05). However, the average yield from the perlite treatment

(8.7 kg/m2 with a corresponding BE of 103%) was highest among the various treatments

(Table 4.1). Hence, perlite was adopted as the substratum for all subsequent treatments.

Table 4.1. Effect of different water-holding materials used as a single layer substratum on mushroom yield and bioefficiency Water-holding Yield Bioefficiency Material Treatment1,2 (kg/m2) (%) Perlite 8.68 ± 1.16a 102.9 ± 13.7a Vermiculite 7.52 ± 1.62a 89.2 ± 19.2a Polyacrylamide 6.82 ± 1.88a 80.9 ± 22.3a Sand 6.65 ± 1.18a 78.8 ± 14.0a Polyacrylamide + perlite 6.13 ± 0.87a 72.7 ± 10.3a 1 Data presented with mean ± standard deviation 2 Treatments followed by similar letters are not significantly different (p>0.05)

Table 4.2 summarizes the results of treatments cased with non-sterile casing in

which S41 rate, perlite volume, and casing type were varied. All main effects were significant for yield and BE (p<0.05), and two of the two-way interactions (rate x perlite

and rate x casing) were significant for yield (p<0.05). All of the two-way interactions for

BE were significant (p<0.05). The three-way interaction (S41 rate x perlite volume x

casing type) for yield and BE was not significant (p>0.05).

61 Table 4.2. Effect of adding perlite underneath a non-composted substrate of commercial grain spawn supplemented with S41 and cased with a sterilized casing containing 25% activated carbon or non-sterile casing on mushroom yield and substrate bioefficiency Perlite Supplement Yield1 Bioefficiency1 Casing Type (ml) (%) (kg/m2) (%) Sterile AC 3000 0 8.25 ± 1.23 107.6 ± 15.9 Sterile AC 3000 5 12.15 ± 0.96 150.9 ± 11.5 Sterile AC 3000 10 13.92 ± 1.84 165.0 ± 21.8 Sterile AC 3000 15 13.51 ± 2.21 153.2 ± 25.1 Sterile AC 3000 20 13.18 ± 2.57 143.3 ± 27.9 Sterile AC 2000 0 8.07 ± 1.62 105.2 ± 21.1 Sterile AC 2000 5 14.28 ± 0.73 177.4 ± 9.1 Sterile AC 2000 10 13.96 ± 1.11 165.5 ±13.2 Sterile AC 2000 15 13.28 ± 2.75 150.7 ± 31.2 Sterile AC 2000 20 13.91 ± 1.00 151.2 ± 10.9 Sterile AC 1000 0 9.07 ± 1.06 118.3 ± 13.8 Sterile AC 1000 5 13.08 ± 2.08 162.5 ±25. Sterile AC 1000 10 13.39 ± 3.61 158.8 ± 42.9 Sterile AC 1000 15 10.50 ± 2.57 119.2 ± 29.3 Sterile AC 1000 20 14.07 ± 2.48 153.0 ± 26.9 Sterile AC 0 0 8.77 ± 1.15 114.5 ± 14.9 Sterile AC 0 5 9.61 ± 1.94 119.5 ± 24.1 Sterile AC 0 10 7.4 ± 0.68 87.8 ± 8.1 Sterile AC 0 15 7.19 ± 0.76 81.6 ± 8.7 Sterile AC 0 20 8.82 ± 1.99 95.9 ± 21.6 Non-sterile 3000 0 0.00 ± 0.00 0.00 ± 0.00 Non-sterile 3000 5 6.20 ± 0.96 77.1 ± 19.4 Non-sterile 3000 10 7.93 ± 1.22 94.0 ± 21.6 Non-sterile 3000 15 10.22 ± 0.39 115.9 ± 7.3 Non-sterile 3000 20 9.61 ± 0.65 104.5 ± 11.4 Non-sterile 2000 0 0.00 ± 0.00 0.00 ± 0.00 Non-sterile 2000 5 9.94 ± 1.95 123.4 ± 39.4 Non-sterile 2000 10 8.68 ± 0.71 102.9 ± 13.7 Non-sterile 2000 15 7.61 ± 0.56 86.3 ± 10.3 Non-sterile 2000 20 11.49 ± 1.42 124.9 ± 25.0 Non-sterile 1000 0 0.00 ± 0.00 0.00 ± 0.00 Non-sterile 1000 5 8.54 ±1.61 106.1 ± 32.5 Non-sterile 1000 10 8.95 ± 0.85 106.2 ± 16.3 Non-sterile 1000 15 9.72 ± 0.123 110.2 ± 2.3 Non-sterile 1000 20 13.57 ± 2.51 147.6 ± 27.4 Non-sterile 0 0 2.11 ± 0.91 27.5 ± 11.8 Non-sterile 0 5 6.19 ± 0.47 76.9 ± 5.8 Non-sterile 0 10 4.79 ± 0.61 56.7 ± 7.2 Non-sterile 0 15 4.32 ± 1.89 48.9 ± 21.5 Non-sterile 0 20 5.76 ± 0.26 62.7 ± 2.9 1 Data presented with mean ± standard deviation

62 Regarding the main effects, the rate of S41 markedly increased mushroom yield

and BE for treatments with 5% S41 compared to treatments with 0% S41, and a marginal

decrease followed by an increase in yield and BE was detected for S41 rates ranging from

5% to 20%. Differences in yield and BE for the 5, 10, 15 and 20% S41 rate were not

significantly different (p>0.05). The effect of perlite volume on mushroom yield was, on

average, highest for the 1000 and 2000 ml perlite treatments, whereas BE peaked for the

2000 ml treatments. Mushroom yield and BE were also was higher with the sterilized

than non-sterilized casing. Figure 4.1 and 4.2 depict the change in mushroom yield and

BE with respect to the different main effects. ) 2 Yield (kg/m

Figure 4.1. Summary of the main effects (S41 rate, perlite volume, and casing type) on mushroom yield (kg/m2).

63

128.2 Rate Perlite Cas ing

107.3

86.4

(%) Bioefficiency 65.4

44.5 0 5 0 5 0 0 0 0 0 le le 1 1 2 0 0 0 i i 10 20 30 ter ter -S S n No

Figure 4.2. Summary of the main effects (S41 rate, perlite volume, and casing type) on BE (%). Overall, the highest mushroom yield was 14.28 kg/m2, which was observed for the 5% S41 + 2000 ml perlite + sterilized casing (AC) treatment. The BE of this treatment was 177%. The lowest yield of mushrooms (0 kg/m2) was observed for 0% S41

+ volumes of perlite (1000, 2000, and 3000 ml) + non-sterilized casing treatments, whereas for the 0 ml perlite treatment, a low mushroom yield of 2.11 kg/m2 (BE =

27.5%) was also observed.

For treatments cased with a sterilized casing (AC), the highest yield among the

0% S41 treatment level was observed for the 1000-ml perlite treatment, which yielded

9.07 kg/m2 with a corresponding BE of 118%. For the non-sterile casing treatments, the highest yield among the 0% S41 treatment level was observerd for the 0-ml perlite, which

64 yielded 2.11 kg/m2 with a corresponding BE of 27.5%. Treatment means among the 0%

S41 rates + at all perlite volumes + sterilized casing with AC were not significantly

different (p>0.05). Increasing the rate of S41 significantly improved yield compared to the 0% S41 treatments (p<0.05). For treatments with both non-zero S41 and perlite levels, the highest yield for the 5% and 10% S41 rates was observed for the 2000 ml perlite treatment, yielding 14.28 kg/m2 and 13.96 kg/m2, respectively. The highest yield

for 15% and 20% S41 was observed for 3000 ml (12.49 kg/m2) and 1000 ml (13.01 kg/m2) perlite.

For treatments receiving non-sterilized casing, mushrooms failed to form when

S41 was not added. However, the 0 ml perlite treatment with 0% S41 yielded mushrooms. However, mushroom yield for the latter was low compared to sterile casing

(AC) treatments. For the 5% S41 treatments, the highest yield was observed for 2000-ml perlite treatment (9.94 kg/m2; BE = 123%). The highest yield among the 10% S41

treatments was observed in the 1000-ml perlite treatment (8.95 kg/m2; BE = 106.2%)

whereas the highest yield among the 15% S41 treatments was observed for the 3000-ml

perlite treatment (10.22 kg/m2; BE = 115.9%). Finally, the highest yield among the 20%

S41 treatments was observed for the 1000-ml perlite treatment (13.57 kg/m2; BE =

148%). Table 4.2 summarizes the results of treatments cased with non-sterilized casing in

which S41 rate and perlite volume were varied. Figure 4.3 provides a graphical summary

of the interaction effects on yield for the three different factors.

65 Yield (kg/m 2 )

Figure 4.3. Summary of significant interaction effects of S41 rate, perlite volume, and casing type on mushroom yield (kg/m2).

As shown in Table 4.3, there was no significant difference in mushroom yield between substrates containing the rye grain and millet grain spawns (p>0.05). For each spawn type, there was an increase in yield with an increase in the rate of S41. However, the differences among the treatment means beyond the 5% S41 rate were not significantly different (p>0.05). The highest yield was observed for the 15% S41 + millet grain spawn treatment (12.03 kg/m2; BE = 136%).

66 Table 4.3. Comparison between millet grain spawn and rye grain spawn on mushroom yield and substrate bioefficiency Rate of Nutrient Yield Grain Spawn Bioefficiency Supplement Yield Type1,2 (%) (%) (kg/m2) Millet 0 3.43 ± 0.92a 44.7 ± 11.9a Millet 5 9.27 ± 0.63b 115.2 ± 7.8b Millet 10 11.08 ± 0.53b 131.4 ± 6.0b Millet 15 12.03 ± 2.39b 136.4 ± 27.1b Millet 20 11.22 ± 0.59b 121.9 ± 6.5b Rye 0 2.47 ± 1.15a 32.3 ± 14.9a Rye 5 11.41 ± 0.32b 141.7 ± 4.0b Rye 10 9.38 ± 1.53b 111.2 ± 17.3b Rye 15 10.41 ± 1.39b 118.1 ± 15.9b Rye 20 10.69 ± 0.38b 116.2 ± 4.1b 1 Data presented with mean ± standard deviation 2 Treatments followed by similar letters are not significantly different (p>0.05)

4.5. Discussion

In accordance with previous findings (Bechara et al., 2005), the addition of water-

holding materials beneath a non-composted commercial grain spawn + S41 substrate

increased mushroom yield compared to treatments without any water-holding materials.

There were no significant differences among the different types of water-holding

materials. Hence, one can opt for the cheapest water-holding material to prevent water-

logging conditions at the tray/substrate interface. One can argue that the use of water-

holding materials might not be necessary if drainage holes are made in the trays.

However, the role of a water-holding material as a water reservoir would be lost. The

commercial grain spawn substrate layers used were relatively thin compared to typical

commercial composted substrates, making substrate drying of great concern. On

average, perlite produced the highest yield of mushrooms and was used for all other

treatments.

67 Several observations can be made for treatments in which the S41 rate, perlite

volume, and casing type were varied. Perlite addition below the substrate up to the 1000-

2000 ml increased mushroom yield. Overall, the use of sterilized casing with activated

carbon improved mushroom yield compared to non-sterilized casing. However, with increasing rates of S41, mushroom yield from the non-sterilized casing treatments approached that obtained from the sterilized casing treatments. Considering that S41 is coated with a proprietary antimicrobial, the increased rates of S41 possibly inactivated microbial contaminants originating from the non-sterile casing material. Thus, it appears

as though the casing layer is limiting mushroom production. This could be due to resident

microbes in the casing that hinder or compete with A. bisporus. This observation could

also be due to a mushroom yield enhancing effect of activated carbon. Overall, the 2000

ml perlite produced the highest BE and is recommended. More work is needed to address

these observations.

In contrast to the findings of Fritsche (1988) and San Antonio (1971), there was

no significant difference between millet and rye grain spawns on mushroom yield

(p>0.05). Hence, if a commercial grain spawn substrate is used for commercial

mushroom production, exchanging between commercial rye grain spawn and millet grain

spawn would have no effect on mushroom yield. Market price and availability of the

grains should control the choice.

4.6. Conclusions

Different factors were evaluated for their effect on mushroom yield and substrate

bioefficiency using substrates composed of commercial grain spawn, S41 supplement and

an underlying layer of water-holding materials. Yield of mushrooms was not affected by

68 the type of water-holding material placed beneath the substrate. However, on average,

perlite produced the highest mushroom yield (8.68 kg/m2) and was used in all additional

testing. S41 rate, perlite volume, and casing type (sterilized or non-sterilized with an activated carbon amendment) were evaluated for their effect on mushroom productivity.

Overall, all three factors had a significant effect on the yield of mushrooms (p<0.05). On

average, yields of treatments with non-sterilized casing were lower than with sterilized

casing with AC. Furthermore, the effect of adding perlite (1000, 2000, and 3000 ml) below the substrate increased mushroom yield. Finally, there was no significant

difference in mushroom yield between rye and millet grain spawns as the basal substrate

material (p>0.05). The highest yield among all treatments of 14.28 kg/m2 and a

corresponding BE of 177% was observed the 5% S41 + 2000 ml perlite + sterilized

casing with activated treatment.

Acknowledgments

The authors would like to acknowledge the assistance of Tom Rhodes, manager,

and the entire crew of the Penn State mushroom research facilities. Funding for this study

was provided by the College of Agricultural Sciences and the Department of Agricultural

and Biological Engineering (Penn State). The grain spawn was generously supplied by

Mark Spear from Sylvan Spawn Inc.

69 Chapter 5

Effect of Delayed-release Supplements in Grain-based Substrate on Yield of the Mushroom (Agaricus bisporus)

5.1. Abstract

Non-composted grain-based substrates mixed with delayed-release supplements

were used as a mushroom substrate to test suitability for mushroom production as a

replacement for the environmentally problematic composted substrate. Five delayed-

release supplements (S41, S44, Promycel Target, T6 and T7) were added at seven rates

(0, 5, 10, 15, 20, 25 and 30%) to a non-composted substrate composed of commercial

millet grain spawn. Overall, mushroom yield, substrate bioefficiency, and mean

mushroom weight increased with the addition of the supplements when compared to the

0% supplement treatment. The highest mushroom yield and substrate bioefficiency were observed for the 20% S41 treatment, which produced 13.73 kg/m2 of mushrooms with a

corresponding substrate bioefficiency of 133.7%. The observed yield and substrate

bioefficiency were considerably higher for S41 than Promycel Target, S44, T6, and T7.

Mean mushroom weight was highest for the 25% T7 treatment (30.2 g), and higher mean

mushroom weights corresponded to treatments with lower mushroom yield. Substrate

temperatures for the different treatments were either similar or lower than the ambient air

temperature. Hence, substrate overheating as encountered in compost-based substrate

was ruled out as being the reason underlying the lower yields observed at the higher rates

of supplementation.

Keywords: Solid-state fermentation, non-composted substrate, alternative mushroom

production systems.

70 5.2. Introduction

Commercial production of Agaricus bisporus (J. E. Lange) Imbach consists of

growing the mushroom on a partially composted organic matrix (Schisler, 1982).

Although widely used, the composting step within the mushroom production process is increasingly viewed as a nuisance, because of malodorous volatile emissions, nutrient-

rich run-off, and accumulation of spent substrate (Heinemann et al., 2003; Heinemann, et

al., 2004). As a consequence of these environmental problems, work on non-composted

mushroom substrates, although not new, has regained momentum (Till, 1962; San

Antonio, 1971; Mee, 1978; Sanchez and Royse, 2001; Mamiro et al., 2007). San Antonio

(1971) developed a non-composted substrate composed of cased rye grain spawn, the

vehicle used to inoculate composted substrates, and concluded that mushroom yield was

comparable to yield from composted substrates. Bechara et al., (2006a) adapted the non-

composted substrate formulation used by San Antonio for a substrate composed of millet

grain spawn. Yield from cased millet grain spawn (8.7 kg/m2) was comparable to

composted substrates (7.7 kg/m2).

In traditional composted substrates, mushroom yield has increased markedly

(average ~ 28.9 kg/m2 – USDA, 2005-2006) due to the development of high-yielding

hybrid strains, improvements in the composting process and, most importantly, the

addition of nitrogen-rich compounds (Sonnenberg, 2000). In Agaricus mushroom

production, nitrogen-rich compounds are classified either as activators or as supplements

(Nair et al., 1993). The term “compost activator” refers to N-rich compounds that are

added to the organic mix with the goal of hastening the composting process (i.e.

increasing the temperature surge) and incorporating the nitrogenous compounds into the

71 thermophilic biomass (Nair et al., 1993). Supplements are materials, typically high-N compounds, added to the mushroom substrate after the composting phase to enhance vegetative growth of A. bisporus and ultimately increase mushroom yield (Gerrits, 1988;

Nair et al., 1993). However, adding high-N compounds to mushroom substrate increases

microbial activity, which increases compost temperature and can lead to total crop loss

(Schisler and Sinden, 1962; Carroll and Schisler, 1974; Nair et al., 1993). To minimize increases in temperature, delayed-release supplements are used, whereby nutrients are

progressively made available to the mushroom fungus and not to microbial contaminants.

Increased mushroom yields of 10-60% were observed when composted substrates were

supplemented with high-N containing delayed-release supplements at spawning (Carroll

and Schisler, 1976). Some methods of creating delayed-release supplements are

formaldehyde-based coatings, encapsulation techniques using hydrophobic compounds

containing a fungicide such as thiabendazol, or calcium sulfate coatings (Nair et al.,

1993; Romaine and Marlowe, 1993). Nair et al. (1993) showed that adding high-N

supplements to mushroom casing, as a means to overcome heat surges in mushroom

substrate, is possible. Romaine and Marlowe (1993) developed an intact canola

(rapeseed) seed-based, delayed-release nutrient supplement for Agaricus grown on

compost. The canola embryo was inactivated using heat treatment (drying oven at 95°C for 24 hrs or autoclave at 121°C for 1.5 hrs) to prevent germination. Mushroom yield from the canola-supplemented compost was either comparable or greater than mushroom

yield from compost supplemented with Spawnmate II SE (a commercially available

supplement). Dahlberg (2004) showed that the addition of purified cellulose to mushroom

substrates increased mushroom yield as well. He concluded that the stimulatory effect on

72 mushroom yield of delayed-release nutrient supplements was due to the carbon portion

and not to the nitrogen portion.

Mushroom casing is a top-dressing composed of various materials (neutralized

peat moss, spent mushroom substrate, soil and others) that is spread over the composted substrate. Typically, mushroom growers refrain from heat-treating their casing material.

However, Bechara et al. (2006a) showed that autoclaved neutralized peat moss casing containing activated carbon (AC) improved the reliability and consistency of small-scale fruiting of A. bisporus. Furthermore, including a layer of perlite, a water-holding material that can carry four times its weight in water, beneath a commercial grain spawn substrate significantly increased mushroom yield from 7.5 kg/m2 to 13 kg/m2 (Bechara et al.,

2005b). It was proposed that the perlite acted as a source and sink of water, whereby excess water from the casing layer accumulates in the perlite reservoir, preventing water- logging conditions and supporting the development of the fruiting bodies. The importance of perlite as a source of water is more plausible when looking at the work of

Kalberer (1990), in which he states that A. bisporus receives 54-83% of its water

requirement for mushroom production from the composted substrate, and 17-46% from

the casing.

The purpose of this study was to evaluate different rates of five commercial

delayed-release nutrient supplements added to non-composted millet grain spawn-based

substrates for their effect on mushroom yield, substrate bioefficiency and mean

mushroom weight.

73 5.3. Methods

5.3.1. Mushroom Grain Spawn Supplements and Water-holding Material

Commercial aged off-white mushroom hybrid millet grain spawn A15 (Sylvan

Spawn Laboratories, Kittanning, PA) with a moisture content of 46% (wet basis) was used in this study. Five delayed-release nutrient supplements, S41, S44, T6, T7 (Full

House- Beta Spawn Co. Inc., Toughkenamon, PA) and Promycel Target (Spawn Mate,

Watsonville, CA) were used. The suppliers describe S41 and S44 as delayed-release supplements composed of roasted, full-fat cracked soybean particles coated with proprietary antimicrobial mixtures, whereas T6 and T7 are a blend of materials that resist surges in compost temperature and green mold. The T7 supplement has a higher protein content than T6. Promycel Target is composed of a blend of proteins (48%), lipids, carbohydrates and micronutrients. All treatments received an underlying layer of horticultural-grade perlite (Therm-o-Rock East, New Eagle, PA).

5.3.2. Preparation of Mushroom Production Containers

Sterilized (121°C for 60 min) polypropylene containers (0.3 x 0.16 x 0.09 m deep) filled with 4.2-cm layer of wetted perlite were used for mushroom production. The ingredients of the mushroom substrate were composed of a basal substrate of commercial millet grain spawn to which a delayed-release supplement was added. Each container was filled with 800 g on a wet weight basis of a mixture of spawn and supplement.

Sterilized casing was prepared by mixing the casing with 25% activated carbon

(Fisher Scientific, Hampton, NH) (v/v) and then autoclaving the mixture for 2 hr (121°C,

103.7 kPa). The containers were transported to a environment-controlled production room.

74 5.3.3. Settings in Mushroom Tray Reactor

All trials, irrespective of treatments, were placed in a mushroom tray reactor at 22

ºC and 80-90% relative humidity (RH). The casing layer was kept moist by watering daily with tap water using a rose-face nozzle. Once the mycelium covered 50%-70% of the casing surface in all mushroom production containers (on average 10 days after placing in chamber), the mycelia was pinned by reducing the temperature to 16 ºC and increasing room ventilation to achieve a carbon dioxide concentration of 800 ppm.

Mushrooms were produced on average 21 days after preparing the mushroom production containers. The mushrooms were harvested manually as soon as the cap diameter reached 2 to 3 cm with the weight and number of the harvested mushrooms recorded immediately after picking. The harvest duration was three weeks for all treatments.

Temperature was recorded using a digital thermometer for 23 days after casing. The thermometer was inserted in the center of each container up to a depth of 3 cm.

5.3.4. Summary of the Experimental Design

A 5 x 7 factorial design within a complete randomized block design (CRD) was used for this study. Five delayed-release supplements (S41, S44, T6, T7 and Promycel

Target) were added at 0%, 5%, 10%, 15%, 20% 25% and 30% of a basal substrate composed of commercial grain spawn.

5.3.5. Parameters Evaluated and Analysis of Data

All treatments were replicated three times. The results of the treatments were presented in terms of yield/unit area (kg/m2), biological efficiency (fresh weight of mushroom/dry weight of substrates), and average mushroom size (fresh weight of mushrooms/number of mushrooms).

75 Mushroom yield, substrate bioefficiency, and mean mushroom weight data were statistically analyzed using the General Linear Model with α = 0.05 for the analysis of variance. For comparisons among treatments, the Multiple Comparison with Best method

(Dunnett Simultaneous Method) for comparisons at α = 0.05 was used. MINITAB

Statistical Software Package (Release 13.1, State College PA) was used for the statistical analysis.

5.4. Results

5.4.1 Mushroom Yield

The effects of the two factors (delayed-release supplement type and rate) including the factor interaction (supplement type x rate) were significant for mushroom yield (p<0.05). The highest mushroom yield (13.73 kg/m2) was observed for the 20% S41 treatment (Table 2), whereas the lowest (0 kg/m2) was observed for all 0% supplement treatments. Overall, mushroom yield was greatest for treatments supplemented with S41, followed by treatments supplemented with Promycel Target, T6, S44 and T7 (Figure 5.1).

76

14 S41 S44 12 Prom

) 10 T6 2 T7 /m

g 8 k ( Figure 1. Mushroom yields for non-composted substrates composed of commercial6 millet grain spawn and different rates of five delayed- Yield (kg/m2) Yield

Yield release4 supplements.

2

0

0 5 10 15 20 25 30 rate (%) Supplement Rate (%)

Figure 5.1. Mushroom yields for non-composted substrates composed of commercial millet grain spawn and different rates of five delayed-release supplements.

The highest mushroom yield for Promycel Target (9.33 kg/m2) was observed for the 15% rate (Table 5.2), whereas yield for the S44 supplement (6.13 kg/m2) was observed for the 10% rate (Table 5.2). For the T6 and T7 supplements, the highest yield was observed for 15% rate (7.44 kg/m2) and 10% rate (6.59 kg/m2), respectively (Table

5.2). In almost all treatments, the increase in mushroom yield for a supplement rate beyond 5% was not significant (p>0.05). For the S44 supplemented treatments, none of the treatments were significantly different (Table 5.2).

77 Table 5.1. Summary of yield, substrate bioefficiency and mean mushroom weight for a basal substrate composed of commercial grain spawn supplemented with S41, S44, Promycel Target, T6 and T7 delayed-release supplements. Mushroom Yield Bioefficiency Mean Mushroom Treatment1,2 (kg/m2) (%) Weight (g) 0% Supplement 0.00 ± 0.00b 0.0 ± 0.0b - 5% S41 9.28 ± 2.16a 99.7 ± 23.2a 11.8 ± 2.8a 10% S41 9.75 ± 2.25a 101.2 ± 23.4a 11.3 ± 2.6a 15% S41 11.81 ± 3.92a 118.7 ± 39.4a 13.3 ± 4.4a 20% S41 13.73 ± 2.04a 133.7 ± 19.9a 11.0 ± 1.6a 25% S41 13.16 ± 2.36a 124.3 ± 22.3a 12.3 ± 2.2a 30% S41 5.17 ± 3.82b 47.5 ± 35.1b 12.2 ± 9.0a 5% S44 5.21 ± 3.48b 55.9 ± 37.3b 19.2 ± 12.8a 10% S44 6.13 ± 2.34b 63.7 ± 24.3b 16.7 ± 6.4a 15% S44 4.95 ± 1.34b 49.8 ± 13.4b 18.8 ± 5.1a 20% S44 5.34 ± 3.64b 52.0 ± 35.5b 12.2 ± 8.3a 25% S44 3.67 ± 1.86b 34.7 ± 17.5b 14.3 ± 7.2a 30% S44 0.72 ± 1.24b 6.6 ± 11.4b 14.7 ± 25.5a 5% Promycel 6.67 ± 4.03b 71.6 ± 43.2b 12.9 ± 7.8a 10% Promycel 8.22 ± 3.99a 85.4 ± 41.4a 14.1 ± 6.8a 15% Promycel 9.33 ± 1.08a 93.8 ± 10.8a 17.9 ± 2.1a 20% Promycel 8.74 ± 2.36a 85.1 ± 22.9a 17.0 ± 4.0a 25% Promycel 9.13 ± 3.88a 86.2 ± 36.6a 16.4 ± 6.9a 30% Promycel 5.89 ± 1.27b 54.0 ± 11.7b 14.4 ± 3.1a 5% T6 6.92 ± 3.75b 74.2 ± 40.3a 16.6 ± 9.0a 10% T6 7.17 ± 1.09b 74.4 ± 11.3a 18.1 ± 2.7a 15% T6 7.44 ± 1.49b 74.8 ± 15.0a 21.0 ± 4.2a 20% T6 6.38 ± 3.32b 62.1 ± 32.3b 34.0 ± 17.7a 25% T6 1.26 ± 1.39b 11.9 ± 13.1b 18.2 ± 19.9a 30% T6 2.81 ± 1.79b 25.8 ± 16.4b 13.1 ± 8.3a 5% T7 5.51 ± 0.18b 59.1 ± 1.9b 26.4 ± 0.9a 10% T7 6.59 ± 4.27b 68.5 ± 44.3b 19.0 ± 12.3a 15% T7 4.17 ± 2.38b 41.9 ± 23.9b 26.1 ± 14.9a 20% T7 0.59 ± 0.83b 5.8 ± 8.1b 28.9 ± 39.7a 25% T7 1.05 ± 0.99b 9.9 ± 9.3b 30.2 ± 28.4a 30% T7 0.63 ± 0.29b 5.8 ± 2.7b 22.8 ± 10.5a 1 Data presented with mean ± standard deviation 2 Treatments followed by similar letters are not significantly different from the best treatment (20% S41 for yield, 20% S41 for bioefficiency and 25% T7 for mean mushroom weight), whereby p<0.05 is significant.

78 5.4.2. Substrate Bioefficiency

The effects of the main factors (supplement type and rate) and their interaction

(supplement type x rate) on substrate bioefficiency were significant (p<0.05). On

average, the highest substrate bioefficiency was observed for S41 and Promycel,

followed by T6, S44 and T7. With the exception of S41, the 10% and 15% supplement

rates had the highest bioefficiency. Aside from the 0% supplement level, the lowest

bioefficiency was observed for all the 30% rates, especially the T7 supplement (Table

5.2). The 20% rate of S41 had a bioefficiency of 133.7% (Table 5.2), whereas the highest bioefficiency for S44 of 63.7% was observed for the 10% rate (Table 5.2). For Promycel,

T6 and T7, the highest observed bioefficiencies were observed for the 15% (93.8%), 15%

(74.8%) and 10% (68.5%) rates, respectively.

5.4.3. Mean Mushroom Weight

For mean mushroom weight, supplement type was the only factor that was significant (p<0.05). The lowest mean mushroom weight was observed for the S41 treatments, whereas the highest was observed for T7; as noted earlier, the highest yielding treatments were observed for the S41 supplement and the lowest yielding treatments were observed for T7. The general trend was an inverse relationship between mean mushroom weight and substrate yield.

5.4.4. Substrate Temperature

Substrate temperatures were rarely over the ambient temperature of the tray reactor. In some cases, especially between day 15 and 21, substrate temperature was lower than ambient temperature, presumably due to a high rate of evapotranspiration.

Temperature dropped to 13 °C on day 23 because of a malfunction with the humidifier.

79 Figure 5.2 shows the substrate temperature readings for the different treatments over 23 days.

23 23

22 22

21 21

20 20 Env Env 19 0% 19 0% 5% S41 10% S41 18 5% S44 18 10% S44 5% Prom 10% Prom 17 5% T6 17 10% T6 5% T7 10% T7 16 16

15 15

14 14

13 13 1 3 5 7 9 11131517192123 1357911131517192123 Day Day

23 23 22 22

21 21

20 20 Env Env 0% 19 0% 19 15% S41 20% S41 18 15% S44 18 20% S44 15% Prom 20% Prom 17 17 15% T6 20% T6 15% T7 16 16 20% T7

15 15

14 14

13 13 1 3 5 7 9 11 13 15 17 19 21 23 1 3 5 7 9 11131517192123 Day Day

23 23

22 22

21 21

20 20 Env Env 0% 19 0% 19 25% S41 30% S41 18 25% S44 18 30% S44 25% Prom 30% Pro 17 17 25% T6 30% T6 25% T7 16 16 30% T7

15 15 14 14 13 13 1 3 5 7 9 11 13 15 17 19 21 23 1357911131517192123 Day Day Figure 5.2. Comparison of the temperature profiles during the 23-day period after casing for the production room (Env.) and non-composted substrates composed of commercial millet grain and five different five delayed-release supplements.

80 5.5. Discussion

The results showed that the addition of delayed-release supplements, typically

used in commercial compost-based substrates, to a non-composted substrate composed of commercial millet grain spawn increased mushroom yield, substrate bioefficiency and mean mushroom weight. The treatment without supplement addition failed to produce

mushrooms, and this observation was likely attributed to the age of the commercial spawn. In this study, commercial grade (aged spawn) and not “juvenile” spawn was used.

In previous experiments, grain spawn age was shown to be a significant factor that influenced mushroom yield; older grain spawn as sold to mushroom producers was less productive than newly colonized “juvenile” spawn (unpublished data).

S41 was the most suitable supplement for A. bisporus mushroom production using

a basal substrate of commercial millet grain spawn, and this concurs with earlier findings

(Bechara et al., 2005a). In the latter study, S44-supplemented substrates were prone to mold contamination, and substrate temperature was not monitored. In the present study, a

high level of contamination was not observed, but some supplements showed some

molding at the higher rates. The reduced yield with S44 compared to S41was not related

to an excessive substrate temperature (Fig. 5.2), but can only be attributed to differences

in antimicrobial coatings since both supplements are composed of cracked soybean. In a

previous study, cracked soybean stimulated the productivity of A. bisporus to a greater

extent than other types of oilseed supplements, such as safflower and niger (Bechara et

al., 2006b). Therefore, cracked soybean-based delayed-release supplements appear to be

the most promising for increasing mushroom yield. However, antimicrobial coatings and

delayed-release properties are also important factors in whether or not a supplement will

81 be effective. Promycel Target, T6 and T7 supplements were least efficacious in

increasing mushroom yield. It is important to note that these supplements have a

similarly small mesh size. Adding them to the substrate at lower rates is satisfactory.

However, as the rate of addition is increased, pore size within the mushroom substrate

would decrease, and that might have hindered oxygen transfer rate. Royse and Sanchez

(2001) observed a negative correlation between substrate particle size and Lentinula edodes mushroom yield (smaller particle size translated to lower yield), and they

attributed the reduction in yield to oxygen depletion within the substrate. Ghilidyal et al.

(1992) showed that oxygen concentrations at depths of up to 0.24 m for Aspergillus niger grown in trays was not growth limiting, whereas optimal carbon dioxide levels for A. bisporus vegetative growth ranges from 1000 to 5000 ppm (San Antonio and Thomas,

1972), with a maximum threshold of 20,000 ppm beyond which inhibition of growth was encountered (Flegg, 1985).

Overall, mushroom yield for non-composted substrates used in this study was lower than the average industry yield value of 30 kg/m2 from commercial compost-based

substrates. However, the volume of substrate per unit of surface area of the production

bed is markedly higher for commercial operations compared to the non-composted

substrates used in this study. The depth of commercial composted substrates typically ranges from 16-20 cm, whereas that of the non-composted substrates used in the present study were 2.5 cm.

Mushroom substrate bioefficiencies of 80-90% are considered ideal (Schisler,

1982). Bioefficiencies for the non-composted substrates used herein were higher for

82 many treatments. For example, the bioefficiency for the 20% S41 treatment was 133.7%, which is considerably higher than for traditional composted substrates.

Substrate temperatures were usually close to the temperature of the ambient air in

the production room. Flegg (1985) reported that optimal growth of A. bisporus was encountered at 24°C, whereas Huhnke and Sengbusch (1968) reported 27°C. Growth was drastically reduced at temperatures above 28°C and total inhibition occurred at 32°C. The maximal temperatures of the different non-composted substrates examined in the present study never approached the threshold of inhibition. From day 15 to 23, substrate temperatures were markedly lower than the ambient air. This was due to a malfunctioning

humidifier in the production room, as most of the heat loss likely occurred by way of

evaporative cooling, since relative humidity ranged from 45 to 55%. In most mushroom

cultivation processes, relative humidity is maintained at 85% or higher. A 98% relative

humidity limited the effect of evaporative cooling, whereas at lower relative humidity,

evaporative cooling contributed to thermal heat loss (Smiths et al., 1999).

5.6. Conclusions

Adding delayed-release supplements to a basal substrate consisting of commercial

grain spawn increased mushroom yields compared to a substrate composed solely of

grain spawn. Among the five delayed-release supplements tested, S41 produced the

highest yield and substrate bioefficiency. The 20% S41 produced 13.73 kg/m2, with a

substrate bioefficiency of 133.7%. Overall, increasing the rate of supplementation above

5% did not significantly increase yield. Mean mushroom weight was affected by supplement type, whereas substrates with lower yields generally produced larger mushrooms. Substrate temperature was not influenced by supplement type or rate, as

83 most of the treatments were either at or below ambient air temperature of the production room.

Acknowledgments

The authors would like to acknowledge the assistance of manager Tom Rhodes and the entire crew at the Penn State Mushroom Research Center. Funding for this study was provided by the Penn State College of Agricultural Sciences and the Department of

Agricultural and Biological Engineering. The grain spawn was generously supplied by

Mark Spear from Sylvan Spawn, Inc.

84

Chapter 6

Evaluating the Addition of Activated Carbon to Heat Treated Mushroom Casing for Grain-based and Compost-based Substrates

6.1. Abstract

Two substrates, a non-composted grain spawn substrate and a traditional

composted substrate, each covered with peat-based casing that contained varying

amounts of activated carbon (AC) and each receiving different heat treatment durations,

were tested for Agaricus bisporus mushroom production. The amounts of AC were 0, 5,

10, 15, and 20% v/v, and the heat treatments were 0, 60, and 180 min at 121°C and 103.4 kPa. Overall, the addition of AC up to 10-15% of casing for a grain spawn substrate increased mushroom yield. However, the addition of AC to the casing for compost-based substrates had no significant effect on yield, whereas heat-treating the casing increased yield. The onset of fruiting was retarded in grain spawn treatments not receiving AC with heat treatment durations of 60 and 180 min, whereas this effect was not as apparent for the compost substrates. On average, mushroom yield was greater for the grain spawn substrate (7.6 kg/m2) than for compost substrate (5.9 kg/m2). For grain spawn substrate,

the results show that the addition of AC ranging from 5-10% was adequate for maximum

mushroom production.

Keywords: non-composted substrates; grain spawn; Agaricus bisporus

85 6.2. Introduction

Agaricus bisporus, the common cultivated button mushroom, is traditionally grown on partially composted substrates composed of plant and animal organic matter.

Once the substrate is fully colonized with the mushroom fungus, a layer of lime- neutralized peat moss, termed “mushroom casing”, is added over the substrate. The casing layer can be added before the substrate is fully colonized, resulting in earlier mushroom production (Flegg, 1985). Although widely used, there are several alternatives to peat-based casing, such as soil (clay-loam), vermiculite, and weathered mushroom compost, among others (Schisler, 1982). The addition of casing is one of two major requirements for the production of fruit bodies of many Agaric mushrooms. The second requirement is the alteration of environmental conditions within the mushroom production chamber conducive to fruiting, i.e. temperature reduction and increased ventilation to reduce the level of carbon dioxide evolved from the fungal biomass (Long and Jacobs 1974; Schisler, 1982).

The role of the casing layer in mushroom fruiting has long been debated. First and foremost, the casing provides the support and storage of the water for developing mushrooms and protects the substrate from desiccation (Flegg, 1985). However, not all materials that provide such conditions can be considered as adequate casing. One hypothesis suggests that microorganisms within the casing, specifically Pseudomonas spp. bacteria, are responsible for fruiting, since sterilizing the casing inhibits or retards the formation of mushrooms (Cresswell and Hayes, 1979; Eger, 1972; Hayes et al.,

1969). Furthermore, bacteria were shown to grow either in close proximity or attached to the mycelium through filamentous networks (Masaphy et al., 1987). Other studies

86 suggest that the casing layer forms a gradient of carbon dioxide or other unidentifiable volatile necessary for fruiting (Schisler, 1957; Tschierpe, 1959).

Activated carbon (AC) was shown to consistently reestablish mushroom production in substrates cased with sterilized casing (Bechara et al., 2006b; Long and

Jacobs, 1974; Nobel et al., 2003; Peerally, 1978). Other materials that favor primordia

(precursor of mushrooms) formation in sterilized casing are zeolite, lignite, and charcoal

(Nobel et al., 2003). Verbeke and Overstyns (1991) developed a theory describing the role of activated carbon in heat-treated peat-chalk casing in which AC appears to affect carbon dioxide reactions in the casing water. In non-sterilized casing, the carbon dioxide evolved from the metabolic activity of the mushroom fungus comes in contact with water in the casing and is converted into carbonic acid, which in turn reacts with calcium carbonate to release calcium and bicarbonate ions (Verbeke and Overstyns, 1991). The calcium precipitates oxalate, an iron-chelating compound exuded by the mycelium and having an inhibitory effect on mushroom production, and renders the oxalate non-reactive

(Verbeke and Overstyns, 1991). Another observation made by Masaphy et al. (1987) was that calcium oxalate crystals typically found on mycelium in compost were absent in casing mycelium prior to primordia formation.

When peat-chalk casing is heat-treated, a reduction in the bicarbonate-carbon dioxide buffering reaction occurs, which may lead to less calcium available for oxalate precipitation (Hayes, 1981; Verbeke and Overstyns, 1991). Some activated carbons can absorb carbon dioxide (Cao and Wu, 2005). Furthermore, iron in the reduced form is inhibitory to mushroom production. Oxalate, as stated earlier, is an iron chelator, whereas activated carbon was shown to be capable of absorbing iron compounds with a

87 preference for the Fe2+ ion, which is more inhibitory than Fe 3+ to mushroom primordia formation (Verbeke and Overstyns, 1991; Uchida et al., 2000).

In industry, pasteurization of casing is recommended to reduce the incidence of mushroom pests that are especially prevalent in soil-based and mushroom compost casing

(Schisler, 1982). However, if the pasteurization temperature is prolonged, a delay and reduction in mushroom yield are observed. Furthermore, Schisler (1982) showed that prolonged heat treatment of casing soils increased spore germination of Verticillium fungicola, a mushroom pathogen, artificially introduced to the casing after the heat treatment. Chikthimmah (2006) also showed that prolonged heat-treated casing was conducive to artificial re-introduction and growth of Listeria monocytogenes and

Salmonella spp., whereas these pathogens were inhibited when introduced to non-sterile casing.

In our earlier studies, we developed a non-composted substrate consisting in part of commercial grain spawn mixed with various amounts of perlite (Bechara et al., 2006a).

Additionally, it was shown that adding a nutrient supplement typically used for composted substrates increased mushroom yield (Bechara et al., 2005a). Furthermore, yield of mushrooms was increased from 7.5 kg/m2 to 13 kg/m2 when the substrate was added over a 4 cm layer of wetted perlite (2 L volume).

The effect of sterilized casing on mushroom production has been studied mostly on composted substrates or sterilized composted substrates, and less on non-composted grain substrates. Furthermore, there is no significant literature to date that explores different concentrations of AC in casing and its effect on mushroom yield. Therefore, the primary objective of this study was to compare the effect of AC application rate, heat

88 treatment duration, and substrate type for peat-based casing on mushroom yield, average

mushroom size, and onset of fruiting. The secondary purpose was to formulate a casing

treatment that could be used for laboratory-scale trials of non-composted substrate, to lessen variability induced by contaminants present in the casing material.

6.3. Methods

6.3.1. Spawn, Nutrient Supplement and Substrate

Off-white aged hybrid millet grain spawn (Amycel Maxx, Avondale, PA) was

used in this study. The grain spawn was stored at 5 °C until use. Composted substrate

was obtained from the Mushroom Research Center at The Pennsylvania State University.

A delayed-release nutrient supplement (S41, Full House- Beta Spawn Co. Inc.,

Toughkenamon, PA), was used at a rate of 5% of the total weight of the substrate (grain

spawn or compost). The supplement is cracked soybean coated with a proprietary

antimicrobial film.

6.3.2. Casing Preparation

The peat-based casing, primarily composed of wetted black peat and calcium

carbonate, was obtained from the Mushroom Research Center at The Pennsylvania State

University. The casing mixed with the treatment-specified amount of AC (UU 50X200-

Calgon, Columbus, OH) was added to polypropylene containers (L = 0.37 m x W = 0.25

m x D = 0.14 m). The volume added to the containers was 8 L/container. The containers

were covered with aluminum foil and autoclaved at 121 °C and 103.4 kPa for the

treatment-specified time (0, 60, and 180 min). The pH of the casing before and after

autoclaving was measured by taking a 10 g sample and adding 100 ml of de-ionized

water. To measure the decrease in pH induced by heat treating peat moss alone, 100 g of

89 wetted peat moss was placed in flasks and autoclaved for 20, 60, 120, and 180 min at 121

°C and 103.4 kPa. A sample of the peat moss before and after autoclaving was taken for

pH measurement as described above.

6.3.3. Preparation of Mushroom Trays

Autoclavable polypropylene containers (L = 0.3 m x W = 0.16 m x D = 0.09 m) were used as mushroom production trays. The trays were filled with 2000 ml of wetted perlite (Therm-o-Rock, New Eagle, PA) giving a depth of approximately 0.042 m, covered with aluminum foil and sterilized for 20 min (121°C, 103.4 kPa). Where grain spawn was used as the substrate, the grain spawn and 5% S41 were mixed and layered over the perlite in the containers (760 g of grain spawn and 40 g of S41/container). The casing was added over the grain spawn substrate. Trays containing composted substrate were prepared by adding 760 g of compost and 40 g of S41 supplement. Casing was added immediately over the composted substrate in the trays. Weights given are wet weights. The moisture content of the grain spawn was 40% (wb), whereas it was 72%

(wb) for the compost substrate (as determined by oven drying at 105 °C for 48 hrs). All trays were transported to a tray bioreactor at the Mushroom Research Center.

6.3.4. Conditions in Tray Bioreactor

Agaricus bisporus trays were placed in a tray bioreactor (4.9 m x 3 m) with

ambient temperature controlled at 22 °C. The trays were watered on a daily basis using a

rose-face nozzle to keep the casing layer moist. Once the mycelium emerged from the

peat moss, conditions in the tray bioreactor were altered to induce fruit body production.

For A. bisporus, this consisted of decreasing the temperature to 16 °C and increasing

ventilation to reduce carbon dioxide concentrations below 1200 ppm. Carbon dioxide,

90 relative humidity, and temperature were monitored regularly. A carbon dioxide measuring device (Telair 7001, Goleta, CA, 93117) was used and the procedure for taking measurements consisted of placing the instrument in the tray bioreactor for a minimum of 15 min or until the carbon dioxide readings stabilized. Relative humidity and temperature were recorded using a hygrometer with a temperature sensor (Extech, model

444712, Extech Instruments, Waltham, MA). Once mushrooms started forming, they were harvested and their weight and number were recorded immediately after harvest.

6.3.5. CO2 Absorption Test

A manometric method using Oxitop® system (WTW, Giessen, Germany) was

used for testing carbon dioxide absorption by AC. The system is composed of a 1.14 L

glass bottle to be partially filled with the biomass sample and a bottle sealing cap

composed of a receptacle for a carbon dioxide absorbent (typically NaOH) and a pressure

sensor that transmits the bottle atmospheric pressure reading to a controller. The oxygen

uptake rate can be calculated based on the pressure drop in the apparatus. The pressure

drop is induced by the oxygen consumption by the mushroom fungus. In this study, the

Oxitop® system was used to compare the abilities of AC and NaOH to detect carbon

dioxide absorption capabilities. The Oxitop® bottles were prepared by adding a sample

of 50 g of commercial grain spawn (Amycel Maxx, Avondale, PA). The bottles were

sealed after 4.5 g of NaOH or AC was added to the CO2 adsorbent receptacle in the bottle

cap. The samples were incubated at 24.5 °C for 48 hrs and the change in pressure was

recorded. Each treatment was replicated twice.

91 6.3.6. Experimental Design

A randomized factorial design was executed with AC rate, heat treatment duration, and substrate composition as variables. The specific levels for the AC rate were

0, 5, 10, 15, and 20%, whereas the heat treatment duration was 0, 60, and 180 min. The substrates used were commercial millet grain spawn and composted mushroom substrate, each supplemented with 5% S41. Table 6.1 provides a summary of the treatments. The response variables recorded and calculated were mushroom yield, time to onset of

fruiting, average mushroom size, and substrate bioefficiency. Onset of fruiting is defined

as the number of days from tray preparation until the first harvest. Average mushroom

size was calculated by dividing the total mushroom yield (fresh weight) by the number of

mushrooms picked, whereas substrate bioefficiency was calculated by dividing the total mushroom yield (fresh weight) by the oven-dry weight of the substrate.

Table 6.1. Summary of treatments for both commercial grain spawn-based substrate and compost-based substrate.

Treatment Activated Carbon (%) Heat Treatment (min) 1 0 0 2 5 0 3 10 0 4 15 0 5 20 0 6 0 60 7 5 60 8 10 60 9 15 60 10 20 60 11 0 180 12 5 180 13 10 180 14 15 180 15 20 180

92 The response variables were statistically analyzed using the General Linear Model

(α = 0.05) with MINITAB statistical software (release 13.1). All pairwise comparisons of treatments were assessed using the Least Significant Difference method.

6.4. Results

The pH was reduced for all heat treatment times for peat moss with no added calcium carbonate. However, the highest reduction in pH was 0.32 observed in the 120 min and 180 min autoclaving treatments. This decrease in pH was not detected in heat- treated casing, because of the buffering effect of calcium carbonate. Fig. 6.1 depicts the decrease in pH of peat moss heat-treated for various time intervals in the autoclave.

8.4

8.2

8

7.8

s 7.6

7.4

pH value 7.2

7

6.8

6.6

6.4 0 5 10 15 20 Activated Carbon (% )

Figure 6.1. Change in pH of peat moss heat-treated for different time intervals in an autoclave.

93 When AC was added to the casing material (peat moss plus calcium carbonate),

an increase in pH was observed compared to the same casing without AC. Without AC

addition, the pH was found to be 7.2, whereas the pH of the casing gradually increased with increasing rates of AC, reaching 8 for the 20% AC treatment. There was no significant change in pH before and after autoclaving for all the casing treatments

(p>0.05). Figure 6.2 depicts the increase in pH for the different casing treatments.

3.50

3.40

3.30

3.20

3.10 before 3.00 after 2.90 pH of Peat Moss of Peat pH

2.80

2.70

2.60

2.50 20 60 120 180 Autoclaving Time (min)

Figure 6.2 pH for different casing treatments, before and after autoclaving.

Overall, the yield of mushrooms from the grain spawn substrate treatments was

greater than compost, with the highest yield of 7.6 kg/m2 produced by the grain spawn

substrate cased with 10% AC/0 min heat treatment. Furthermore, the average mushroom

size was greater for the grain spawn substrate compared to the compost substrate.

Regarding substrate bioefficiency, the values for the composted substrate were, on

94 average, higher than the grain spawn substrate, because of the low moisture content in the latter. The highest recorded bioefficiency was 115% for the 15% AC/180 min heat treatment with a compost substrate supplemented with 5% S41.

The effect of casing heat treatment time on mushroom yield was not significant

(p>0.05) in the grain spawn substrate treatments. However, the level of AC had an effect on mushroom yield for all three heat treatment levels (p<0.05). For the 0 min heat

treatment, the highest yield of mushrooms was observed in the 10% AC (7.6 kg/m2)

treatment, whereas the highest yield of mushrooms collected for the 15% AC/60 min heat

treatment and 20% AC/180 min heat treatment were7.2 kg/m2 and, 6.8 kg/m2

respectively. With respect to the 0% AC casing treatments, there appeared to be a

decrease in mushroom yield with an increase in heat treatment duration. However, this

observation was not statistically significant (p>0.05). The effect of heat treatment duration, AC rate, and the interaction of the two factors on the onset of fruiting were significant (p<0.05). In treatments without the addition of AC, the onset of fruiting was delayed in response to heat treatment, with the highest being the 180 min heat treatment

(38 days) and lowest being the 0 min heat treatment (29 days). However, for each heat treatment, the onset of fruiting for 5%, 10%, 15%, and 20% rates of AC was not significantly different. There was no significant difference in mushroom size among all the grain spawn substrate treatments (p>0.05). Table 6.2 summarizes the results for the grain spawn substrate with the different casing treatments.

95 Table 6.2. Summary of results for casing treatments with a grain spawn-based substrate supplemented with 5% S41.

Ave. Yield2 Bioefficiency3 Onset of Treatment1 Mushroom2 (kg/m2) (%) Fruiting2 (days) Size (g) 0% AC/0 min 1.6 ± 0.5a,b 16 ± 5 30 ± 9 a 29 ± 2 d,e 5% AC/0 min 6.3 ± 0.2d,e,f 61 ± 2 30 ± 1 a 27 ± 3 c,d,e 10% AC/0 min 7.6 ± 1.5 f 74 ± 15 31 ± 6 a 21 ± 1 a 15% AC/0 min 5.1 ± 0.5 c,d,e,f 50 ± 4 33 ± 3 a 26 ± 3 b,c,d 20% AC/0 min 3.5 ± 2.1 b,c 34 ± 20 36 ± 21 a 25 ± 3 b,c 0% AC/60 min 1.6 ± 0.5 a,b 15 ± 5 32 ± 10 a 30 ± 3 e 5% AC/60 min 5.7 ± 2.0 d,e,f 55 ± 20 34 ± 12 a 24 ± 3 a,b,c 10% AC/60 min 7.2 ± 0.9 e,f 70 ± 9 33 ± 4 a 23 ± 1 a,b 15% AC/60 min 7.2 ± 1.5 e,f 70 ± 15 33 ± 7 a 23 ± 1 a,b 20% AC/60 min 5.1 ± 1.3 c,d,e,f 50 ± 13 34 ± 9 a 23 ± 1 a,b 0% AC/180 min 0.6 ± 0.5 a 5 ± 5 1 ± 1 a 38 ± 3 f 5% AC/180 min 5.6 ± 0.7 d,e,f 56 ± 7 35 ± 4 a 24 ± 3 a,b,c 10% AC/180 min 4.6 ± 1.8 c,d 44 ± 18 26 ± 10 a 24 ± 1 a,b,c 15% AC/180 min 6.6 ± 1.8 d,e 65 ± 18 21 ± 6 a 23 ± 2 a,b 20% AC/180 min 6.8 ± 0.7 e,f 67 ± 7 34 ± 3 a 23 ± 2 a,b 1 Data presented with mean ± standard deviation 2 Treatments followed by similar letters are not significantly different (p>0.05) 3 Bioefficiency has similar significance indicators as for the yield data

Mushroom yield from the compost substrates was lower on average than that from the grain spawn substrate. For compost substrate, the highest mushroom yield of 5.9 kg/m2 was produced from the 15% AC/180 min treatment. The AC rate did not

significantly affect mushroom yield (p>0.05). However, heat treatment duration

significantly affected mushroom yield (p<0.05). Regarding the onset of fruiting, the

effect of AC rate and heat treatment duration was statistically significant (p<0.05). Heat-

treating the substrate, in fact, reduced the time to onset of fruiting, and, in some

treatments, the addition of AC further reduced the time to onset of fruiting. The heat

96 treatment duration and the addition of AC had no effect on average mushroom yield

(p>0.05). Table 6.3 summarizes the results for the partially composted substrates.

Table 6.3. Summary of results for casing treatments with compost-based mushroom substrate supplemented with 5% S41

Ave. Yield Bioefficiency Onset of Treatment1 Mushroom Size (kg/m2)2 (%)3 Fruiting (days)2 (g)2 0% AC/0 min 2.7 ± 0.2 a,b,c,d 52 ± 3 13 ± 1a 29 ± 2 g,h 5% AC/0 min 1.7 ± 0.4 a 33 ± 7 15 ± 3 a 30 ± 1 h 10% AC/0 min 2.1 ± 0.3 a,b 41 ± 6 13 ± 2 a 28 ± 1 f,g,h 15% AC/0 min 2.5 ± 0.8 a,b,c 49 ± 14 11 ± 3 a 26 ± 2 e,f,g 20% AC/0 min 2.2 ± 0.5 a,b 42 ± 10 13 ± 3 a 28 ± 2 f,g,h 0% AC/60 min 4.2 ± 2.0 c,d,e,f 80 ± 39 17 ± 8 a 23 ± 2 c,d,e 5% AC/60 min 3.5 ± 0.6 a,b,c,d 67 ± 11 16 ± 3 a 23 ± 2 b,c,d 10% AC/60 min 5.1 ± 0.6 d,e,f 99 ± 12 12 ± 1 a 19 ± 1 a 15% AC/60 min 4.9 ± 0.8 d,e,f 96 ± 15 13 ± 2 a 21 ± 1 a,b,c 20% AC/60 min 4.9 ± 1.4 d,e,f 95 ± 27 13 ± 4 a 20 ± 2 a,b 0% AC/180 min 3.9 ± 1.7 b,c,d,e 76 ± 33 13 ±6 a 25 ± 3 d,e,f 5% AC/180 min 5.3 ± 1.5 e,f 103 ± 29 13 ± 4 a 21 ± 0 a,b,c 10% AC/180 min 5.6 ± 1.3 e,f 109 ± 26 12 ± 3 a 22 ± 3 b,c,d 15% AC/180 min 5.9 ± 1.3 f 115 ± 26 13 ± 3 a 21 ± 2 a,b,c 20% AC/180 min 5.8 ± 1.2 f 112 ± 24 12 ± 2 a 21 ± 1a,b,c 1 Data presented with mean ± standard deviation 2 Treatments followed by similar letters are not significantly different (p>0.05) 3 Bioefficiency has similar significance indicators as for the yield data

Finally, for the carbon dioxide absorption tests, the results showed that the AC

used in this study is not a carbon dioxide absorbent. This observation can be seen in

Figure 6.3 which presents the results in terms of pressure drop (kPa) against time (hrs).

There was no pressure drop in treatments with AC added as a CO2 absorbent, whereas

with the NaOH a large pressure drop is observed. The rise in pressure from 0- 8 hrs for

some readings is attributed to temperature differences between the lab and the incubator.

97 6.5. Discussion

The effect of heat-sterilizing casing to inhibit or retard mushroom production has

long been established. In this study, this effect was observed for the grain spawn

substrates and not in the composted substrates. Whether this manifestation of mushroom

inhibition is due to the inactivation of organisms necessary for the fruiting of the

mushroom fungus, or changes in the physical and chemical properties of the casing

material, or a combination of both, has not been ruled out. Some work investigated

sterilizing casing using non-thermal methods such as irradiation to eliminate these effects

(Nobel et al., 2003). The observations indicated that non-thermal sterilization of casing

also induced mushroom inhibition. This is most likely due to the inactivation of the

casing biological component that plays a role in mushroom fruiting. However, as in heat-

treated casing material, mushroom primordia formation was re-established when AC,

lignite, or zeolite was added to the casing (Nobel et al., 2003). With heat-treatment, a

reduction in peat moss pH is observed possibly due to the release of humic acids, whereas

it is not detected in lime-neutralized casing, because of the buffering capacity of the calcium carbonate (Verbeke and Overstyns, 1991). Verbeke and Overstyns (1991) hypothesized that the drop in pH reduces the buffering capacity of the casing, which is critical during the evolution of carbon dioxide from the growth of the mushroom fungus.

The addition of AC increased the pH of the casing material as indicated earlier.

Therefore, if the pH reduction and its effect on calcium carbonate buffering capacity is limiting mushroom production, the addition of AC may help to limit pH change, because of the alkaline nature of the AC used or through the absorption of organic acids evolved from the heating process that would induce a pH decrease as shown in Fig. 6.1.

98

Mushroom production is also inhibited by the presence of the reduced form of

iron (Verbeke and Overstyns, 1991). Heat treating the casing might increase the

availability of iron, possibly due to the increased presence of oxalate released by the

mycelium (Verbeke and Overstyn, 1991). AC can absorb a variety of compounds. These

compounds include ferrous/ferric ions and or gases such as carbon dioxide (Cao and Wu,

2005; Uchida et al., 2000). The absorption of iron onto AC removes the inhibitory effect

of iron, whereas the absorption of carbon dioxide reduces the gas loading effect on the

carbonation reactions in the casing, i.e. comparable to the ventilation effect in mushroom

pinning.

In the present work using heat-treated casing, the grain spawn substrate yield of

mushrooms was highly dependent on the addition of AC regardless of heat treatment.

Although not statistically significant, there seemed to be a marked decrease in mushroom

yield between the 0 min heat/0% AC casing and the 180 min heat/0% AC casing

treatments. Contrary to expectation, the addition of AC in the 0 min heat treatments had

an effect on mushroom yield, which indicates that the addition of AC to casing material

of grain spawn substrate will increase yield, regardless of any heat treatment. In contrast,

there was no significant effect of adding AC to compost-based treatments. However,

these results leave open the possibility that fully colonized substrates of both composted

and non-composted origin may be stimulated by the addition of AC to casing. This was

not tested, because in this study the composted substrate was not fully colonized with A. bisporus at casing. The positive effect of AC in casing for grain spawn substrate

treatments could be a manifestation of carbon dioxide or oxalic acid loading, because the

99 grain spawn is fully colonized, whereas the compost is not. AC used in this study did not

absorb carbon dioxide as shown in Fig. 6.3. Hence, the carbon dioxide absorption effect

is ruled out.

5

0 0 4 8 12162024283236404448 )

-5 ActCarb-1 ActCarb-2 NaOH-1 -10 NaOH-2 Pressure Drop (kPa

-15

-20 Time (hrs )

Figure 6.3. Comparison between activated carbon and NaOH as carbon dioxide absorbent.

Heat treating the casing improved the yield of mushrooms for compost treatments.

This could be attributed to limiting the growth of competing microorganisms introduced into the trays by the addition of the non-sterile casing, which subsequently overgrew the mushroom fungus in the un-colonized compost substrate. This was apparent in some of the 0 min heat casing treatments for compost substrate. Heat-treating the casing inactivated those contaminants, resulting in the observed higher yield. Hence, there is some benefit to heat-treating casing before its application onto composted substrates. It is

100 important, however, to state that compost-based substrates are inherently rich with

various microorganisms that establish themselves in the composting phase of the

mushroom production process. The reduced effect of AC in compost treatments may well

be due to the colonization of casing microflora present in the compost, characterized by

its microbiological diversity compared to grain spawn substrate, which is solely

composed of the mushroom fungus. This hypothesis is supported by Samson et al. (1987)

in which they observed an increase in Pseudomonas putida populations, one of the

bacterial species shown to initiate mushroom primordia (Hayes et al., 1969), during the

colonization phase of A. bisporus in the compost substrate. Therefore, if AC does indeed

replace the biological component of casing and the necessary casing microflora are

present in the compost substrate and subsequently colonize the casing material, the

reduced effect of AC in compost substrates can be explained.

It is anticipated that increasing the heat treatment duration would lead to a longer

time to the onset of fruiting, because more of the casing microflora is inactivated and

more organic acids will be produced from the peat moss, which would affect calcium

carbonate buffering capacity. This observation is reflected well in the grain spawn

substrates in which the 0% AC treatments fruited at 29, 30 and 38 days for the 0 min, 60

min, and 180 min heat treatments, respectively. In contrast, the addition of AC markedly

reduced the onset of fruiting, especially for the 180 min heat treatment. The same trend is observed for the compost with the heat- treated treatments having an earlier onset of fruiting.

There was no difference in average mushroom size among casing treatments within similar substrate trials. However, average mushroom weight for grain spawn

101 substrates were greater than for compost substrates. This could be due to the higher

moisture content of the compost substrate (72%-wb) compared to the grain spawn

substrate (40%-wb).

The use of non-heat treated casing material has become a convention in

commercial mushroom production. The effect of contaminants in the mushroom casing

might not pose as much of a problem when fully colonized mushroom substrate is used.

However, there is always the possibility of introducing contaminants through non-sterile casing. Further research is needed to address the specific questions with regard to the fate of mushroom and human pathogens in sterile and non-sterile casing for both composted and non-composted substrates. However, for the purpose of laboratory-scale studies using

non-composed grain-based substrates, heat-treating mushroom casing along with the

addition of AC would eliminate much of the variability induced by the addition of non-

sterile casing.

6.6. Conclusions

The effect of adding AC to heat-treated casing on compost-based substrates and

on grain spawn-based substrates were different. For composted substrates that are not

fully colonized with A. bisporus, the addition of AC had no significant effect on

mushroom yield or size (p>0.05). However, casing heat treatment time did increase

mushroom yield. The yield of mushrooms from grain spawn substrates also was sensitive

to heat treatment duration and, in contrast to compost-based substrate, benefited from the

addition of AC to the casing. The highest yield was found for casing containing 5-10%

AC. Average mushroom size was not affected by heat treatment duration or AC rate,

whereas onset of fruiting was effected. Based on manometric tests, the differences in

102 observations due to AC could not be due to carbon dioxide adsorption by the AC and is likely due to a microbiological or pH effect. However, the results showed that grain

spawn substrates tend to yield greater when 5-10% AC is added to the casing mixture,

whereas compost-based substrates were not affected by the addition of AC to the casing

mixture.

Acknowledgments

The authors would like to acknowledge the assistance of Doug Keith, and Henry

Shawley. The authors would like to thank Dr. Tom Richard for allowing us to use his

lab’s respirometric equipment. Funding was provided by the College of Agricultural

Sciences and the Department of Agricultural and Biological Engineering, The

Pennsylvania State University.

103 Chapter 7

Cultivation of Agaricus bisporus and Agaricus blazei on Substrates Composed of Cereal Grains and Oilseeds

7.1 Abstract

Agaricus bisporus and Agaricus blazei are commercially produced on composted

organic substrates. To test if the environmentally problematic composting process is

necessary, grain substrates were examined for mushroom production. The basal

substrates consisted of sterilized millet-grain, to which various oilseeds (niger, safflower,

and soybean) were added at the rates of 0%, 15%, and 30% of the total substrate

composition. Furthermore, the effect of the commercial delayed-release supplement S41

added to the substrate at 0% and 5% of total composition was tested for its effect on

yield. The addition of oilseeds to the basal substrate is termed Stage I supplementation,

whereas the addition of S41 before casing is termed Stage II supplementation. Mushroom

production was highest (16.9 kg/m2) for A. bisporus when grown in the basal grain-based

substrate supplemented with 15% soybean and 5% S41. For A. blazei, the highest yield

(15.9 kg/m2) was obtained with the addition of only 30% niger to the basal substrate. In

almost all treatments of A. bisporus, an increase in yield when using the S41 supplement was observed. However, this effect was not comparable with A. blazei trials. The

magnitude of the yield increase due to the S41 amendment was dependent on the type

and rate of oilseed used. Additionally, a complete inhibition in the growth of A. blazei

was observed for all treatments involving 30% soybean. The results indicate that A.

bisporus and A. blazei can be cultivated on sterilized mixtures of grain and oilseeds as

substrates, but the two mushroom species have significantly different nutritional

104 requirements for optimal fructification. Finally, a combination of Stage I and Stage II supplementation can be used to maximize yield from non-composted grain-based substrates.

Keywords: Solid-state fermentation, nutraceuticals, non-composted substrates, activated carbon

7.2. Introduction

Agaricus bisporus (button mushroom) is widely cultivated worldwide, with annual sales in the U.S. generating $841 million (USDA, 2005-2006). The price of the button mushroom is on average $2.50/kg fresh weight, which is $25/kg dry weight if one assumes that 90% of the mushroom weight is water (USDA, 2005-2006). The button mushroom is sold primarily as a fresh product with a shelf life of about a week. The mushroom cultivation system has long been studied and perfected using the composting of plant and animal organic matter as the fundamental step in the substrate preparation process. The tremendous increase in mushroom yield was primarily attributed to the optimization of the composting process that began with Sinden and Hauser (1950) who developed the two-phase composting system. In addition, the development and use of protein-rich delayed-release supplements further increased mushroom yield (Carroll and

Schisler, 1974). Romaine and Marlowe (1993) developed and patented an intact oilseed- based, delayed-release nutrient supplement for composted substrates. They showed that heat-treated oilseeds performed just as well as commercial delayed-release supplements.

Other factors have also contributed to the increase in mushroom production, such as the development of high-yielding hybrid strains, a better understanding of mushroom requirements, and better sanitation. On average, yield of the button mushroom on composted substrates (0.16-0.2 m thick) is 30 kg/m2 (USDA, 2005-2006).

105 Agaricus blazei, a tropical mushroom also know as the almond Portobello, is

cultivated in Japan, China, Korea, Thailand, Indonesia, United States, and Brazil (Chang

and Miles, 2004). The cultivation system for this mushroom is similar to A. bisporus

except that higher temperatures are necessary for growth. Like the button mushroom,

composting is an intrinsic step in the substrate preparation process. A. blazei is primarily

grown because of its medicinal properties, such as combating physical and emotional

stress, improving the immune system, inducing tumor cell death, among others (Chang

and Miles, 2004). The active ingredient is thought to be a beta-D-glucan protein complex

(Mizuno, 2000; Chang and Miles, 2004; Filho et al., 2006). Commercially, dietary

supplements of A. blazei are sold in a dried form and they can be derived from mushroom tissue or mycelium grown in bioreactors using liquid fermentation (Filho et al., 2006;

Intabon et al., 2001). The wholesale price of this mushroom in Brazil ranges from $40-

80/kg dried mushroom, whereas retail prices range from $100-$500/kg dried mushroom

(Filho et al., 2006). In Japan and the U. S., the mushroom can be sold up to $600/kg dried mushroom (Filho et al., 2006). Yields of A. blazei on composted substrates are varied.

Iwade and Mizuno (1997) reported yields of 30-50 kg/m2 (production cycle and

harvesting duration not presented), whereas Filho et al. (2006) reported yields of 8-17

kg/m2 for a production cycle of 70-150 days.

The traditional production of mushrooms is characterized as and termed solid

state fermentation (SSF). SSF is a process whereby microorganisms are grown on a solid matrix (organic or inorganic) in the absence of free flowing water (Nagel, et al., 2000;

Pandey et al., 2000). Typically, many Agaric mushrooms are produced in trays filled

with substrate. Trays are moved to chambers in which environmental conditions are

106 controlled. Such chambers are termed “tray bioreactors” (Mitchell et al., 2003).

As stated earlier, both Agaricus bisporus and Agaricus blazei mushrooms are

produced on composted substrates. Composting, the degradation of organic matter by

microorganisms, is an environmentally problematic step in the mushroom production

process; odor generation and nutrient rich run-off are produced (Bechara et al., 2006a,

2006b). Furthermore, the composting step is labor intensive, requires heavy machinery,

and is time consuming. In addition, once mushroom production ceases, the spent

substrate is steamed to inactivate pathogens and then disposed of in different ways,

including open fields where composting activity resumes and again, odor problems and

nutrient-rich run-off are formed.

The use of various sterilized and pasteurized non-composted substrates for A. bisporus mushroom production has been studied previously (Till, 1962; Huhnke and Von

Sengbush, 1968; Sanchez and Royse, 2001; Mamiro et al., 2007). However, the studies

were generally conducted not with the goal of eliminating composting from the

cultivation process, but rather to show that composting is not a necessary prerequisite for the formation of a suitable substrate for A. bisporus. There are several reports of sterilized

substrates that have been successfully used for A. blazei mushroom production (Mizuno,

2000) including a patent by Iwata and Furuya (2002).

Growing the mycelium of the mushroom fungus on sterilized grains is a mainstay

in the mushroom spawn industry. Grain spawn is the vehicle used to inoculate composted

substrates. It is composed of sterilized cereal grains colonized with the mushroom fungus

(Sinden, 1932). Kananen et al. (2000) developed and patented a grain spawn formulation

containing a mixture of cereal grains and commercial supplement mixtures (oilseed and

107 commercial delayed-release supplements). The yield of mushrooms, on average, from a

compost-based substrate inoculated with the spawn-supplement mixture was greater than

a similar substrate inoculated with the traditional rye grain spawn. A method of

producing A. bisporus fruiting bodies directly from rye grain spawn was developed by

San Antonio (1971). Although the yield of mushrooms was not presented, San Antonio

(1971) concluded that the quantity of mushrooms produced was comparable to that

obtained from conventional compost. Bechara et al., (2005b) developed a non-composted substrate using commercial rye grain spawn supplemented with S41, overlain on a 0.042 m layer of perlite, which yielded 13 kg/m2. S41 is a delayed-release supplement composed of cracked soybean and coated with a proprietary antimicrobial formulation that is traditionally added to composted substrates to increase yield. As for A. blazei,

Iwata and Furuya (2002) patented a grain-based formulation that primarily used foxtail millet and other grains as basal substrate.

No literature was found on systematically evaluating various formulations of grain/oilseed substrates for Agaricus species. Based on findings from previous work with

spawn formulations, supplements, and non-composted substrates, the present study was

conducted. The objectives of this study were two-fold. The first was to determine the

effect of adding various oilseeds at different rates to a millet-grain basal substrate, termed

"Stage I supplementation", for A. bisporus and A. blazei mushroom production. The

second was to evaluate whether the addition of S41 supplement after the grain/oilseed

substrate was colonized with the mushroom fungus, termed “Stage II supplementation”,

could further increase mushroom yield.

108 7.3. Materials and Methods

7.3.1. Fungal Strains, Seeds, and Supplements

Agaricus bisporus (MC 459) and Agaricus blazei (WC 837) were obtained as rye grain spawn cultures (400 g of colonized rye grain in 1000-ml wide-mouth Erlenmeyer flasks) from the Mushroom Spawn Lab at The Pennsylvania State University.

Commercial rye grain spawn (Sylvan 140; Sylvan Spawn Lab, Kittanning, PA) of A. bisporus was included as an external control in evaluating the yield of the oilseed substrates. The A. bisporus spawn was stored at 5°C, whereas A. blazei spawn was kept at

22 °C until use. It is noteworthy to mention that A. blazei did not store well at low

temperatures (5 °C). The millet grain and oilseed (niger, soybean, and safflower) were

purchased from a local Agway store (Pleasant Gap, PA). The S41 supplement was

obtained from the Mushroom Test and Demonstration Facility at The Pennsylvania State

University.

7.3.2. Grain Substrate Preparation Process

The substrate preparation process for the two mushrooms was comparable. Millet

grain and de-ionized water was added to a large autoclavable polypropylene container in

a 1 to 1 mass ratio. The mixture was autoclaved for 15 minutes (slow exhaust rate 3.4

kPa/min) and then allowed to cool to room temperature. The grains had a final moisture

content of 0.6 kg/kg wet basis. Unicorn® (Unicorn Imp. & Mfg. Corp., Commerce, TX)

type 10 autoclavable polypropylene bags with filter patches were used as bag-reactors for

the growth of the mushroom fungus on the grain substrate. Millet was added to the bags

along with the appropriate rates of oilseeds (soybean, niger, and safflower). Each bag

contained 1 kg of grains. Following this step, calcium carbonate and calcium sulfate

109 purchased from Fisher Scientific (Hampton, NH) were added in the ratios of 10 g each/

200 g of grains (millet and oilseeds). The materials were mixed and the bags were sealed

with paper clips and sterilized for 3 hrs in an autoclave (121 °C, 103.4 kPa) at slow

exhaust (3.4 kPa/min). The sterilized material was transferred to a laminar flow hood and

allowed to cool overnight. The following day, the substrate was aseptically inoculated

with 10-20 grains of rye grain spawn. The bags were sealed and transferred to a chamber

where the ambient temperature was at 22 °C for A. bisporus and 25 °C for A. blazei to

promote colonization.

7.3.3. Mushroom Production Tray Preparation

Autoclavable polypropylene containers (L = 0.3 m, W = 0.16 m, D = 0.09 m) were used as trays for the production of the two mushrooms. Preparation of the trays was comparable to the method described by Bechara et al. (2005). In summary, the trays were filled with 2000 ml of wetted perlite (Therm-o-Rock, New Eagle, PA), covered with aluminum foil and sterilized for 20 min (121°C, 103.4 kPa). The colonized substrate was added to the containers (800 g/containers). For treatments with S41 (Full House- Beta

Spawn Co. Inc., Toughkenamon, PA), each tray contained 40 g of S41 and 760 g of grain substrate. After adding the substrate, a mixture of sterilized limed peat moss (120 min,

121 °C, and 103.4 kPa) with 10% (v/v) activated carbon (Fisher Scientific, Hampton,

NH) was overlain on top of the substrate up to a thickness of 2 cm. The pH of the casing was recorded before (7.2) and after autoclaving (7.5). The trays were then transported to separate tray bioreactors.

110 7.3.4. Tray Bioreactor Conditions

Agaricus bisporus trays were placed in a tray bioreactor (4.9 m x 3 m) with an

ambient temperature controlled at 22 °C at the Mushroom Research Center (Penn State).

Agaricus blazei trays were transferred to tray bioreactors that were assembled on-site.

The tray bioreactors were designed commercially as indoor greenhouses (Midwest

Quality Glove Inc, Chillicothe, MO) and consisted of metal structures with four shelves that could hold up to five trays. One tray-bioreactor consisted of three shelves rather than four. All three tray bioreactors were covered by clear polyethylene plastic.

The mushroom production trays were watered on a daily basis using a rose-face nozzle to keep the casing layer moist. Once the mycelium emerged from the peat moss, conditions in the tray bioreactors were altered to induce fruit body production. For A. bisporus, this consisted of decreasing the temperature to 16 °C and increasing ventilation

to reduce carbon dioxide concentrations. As for A. blazei, 9 cm slits were made in the

polyethylene covers to improve ventilation. Two slits were made on the top of the

chambers and the rest were made on the front side. Humidified fresh air was also

introduced to each of the tray bio-reactors using an air pump (Gast®, MFG corp., Benton

Harbor, MI) with an air flow rate of 10 L/min. Humidification of the inlet air was

achieved using 10 L Nalgene carboys filled with tap water. Water was added to the

carboy vessels on a weekly basis.

7.3.5. Measured Variables in Tray Bioreactors

Environmental conditions such as carbon dioxide, relative humidity, and

temperature were recorded on a daily basis. A carbon dioxide measuring device (Telair

7001, Goleta, CA) was used by placing the instrument in the respective tray bioreactors

111 for a minimum of 15 min or until the carbon dioxide readings stabilized. Relative humidity and temperature were recorded using a hygrothermometer (Extech, model

444712, Extech Instruments, Waltham, MA). Once mushrooms started forming, they were harvested and their weight and number were recorded immediately after harvest.

7.3.6. Experimental Set-up and Statistical Analysis

The experimental set-up consisted of a factorial design with the following factors: type of oilseed, oilseed rate, and S41 supplementation rate (Table 7.1). Each treatment consisted of three replicates. In addition, a substrate composed in part of commercial grain spawn (Sylvan 140) and 5% S41 was tested and compared to the millet grain substrate with A. bisporus produced on-site.

Table 7.1 provides a summary of the treatments applied for both fungi. Treatment (#) Oilseed Type Oilseed Rate (%) S41 Rate (%) 1 -- 0 0 2 -- 0 5 3 Niger 15 0 4 Niger 15 5 5 Niger 30 0 6 Niger 30 5 7 Safflower 15 0 8 Safflower 15 5 9 Safflower 30 0 10 Safflower 30 5 11 Soybean 15 0 12 Soybean 15 5 13 Soybean 30 0 14 Soybean 30 5

The results of the treatments are presented as the yield per unit area (kg/m2), substrate bioefficiency, and average mushroom weight. Substrate bioefficiency consisted of dividing the fresh weight of mushroom by the initial dry weight of the substrate and was expressed as a percentage. Average mushroom weight was calculated by dividing the total weight of the harvest for a given treatment by the total number of mushrooms

112 produced. The analysis of variance for mushroom yield, substrate bioefficiency, and

average mushroom weight was done using the General Linear Model (α = 0.05) with

MINITAB statistical software (release 13.1). All pairwise treatments were compared using the Tukey multi-comparison test (p<0.05).

7.4. Results

7.4.1. Production Time

The total production times for the two mushroom-producing species were

different since the time interval between sequential harvesting for A. blazei was 8-15

days, compared to 4-7 days for A. bisporus. For A. bisporus, the total production time

was 70 days, whereas the total production time for A. blazei was 98 days. Both mushroom species were kept in the bags for 32 days before being added to the mushroom trays. Substrates lacking oilseeds tended to colonize faster than those with oilseeds.

Agaricus bisporus initiated fruiting in 17 days compared to 21 days for A. blazei. Table

7.2 provides a summary of the time intervals for both mushroom species.

Table 7.2. Summary of duration of the different steps in the mushroom production process A. bisporus A. blazei Time in bags (d) 32 32 Time in trays (d) 17 21 Time harvesting (d) 21 45 Period between harvests (d) 4-7 8-15 Total production time (d) 70 98

7.4.2. Environmental Conditions in Tray Bioreactors

The A. bisporus tray bioreactor maintained the temperature and carbon dioxide levels within the specified parameters throughout the production phase. A few days before pinning, a layer of overgrown mycelium was detected in all trays. This effect, if

113 left untreated, could decrease mushroom yield by rendering the casing hydrophobic and

impermeable to water (Flegg and Wood, 1985). Hence, the casing layer was ruffled

(breaking the overgrown mycelium in the casing) using a small rake. Relative humidity

fluctuated between 75% and 85%. Figure 7.1 depicts the tray bioreactor conditions throughout the mushroom production phase.

2500 25

2000 20

150 0 15 CO2 Yield Temp 10 0 0 10 Temperature (C) CO2 (ppm) & Yield (g) Yield & (ppm) CO2

50 0 5

0 0 1234567891011121314151617181920212223242526272829303132333435363738 Day

Figure 7.1. Environmental conditions and mushroom yield/day for A. bisporus tray bioreactor

The tray bioreactors for A. blazei were not successful in maintaining the specified

environmental conditions. Temperature fluctuated according to ambient conditions.

Ideally, the temperature during the mushroom production phase should be maintained at

28 °C but the temperature fluctuated between 22 °C and 28 °C. However, there are no

reported negative physiological effects of major temperature fluctuation for A. blazei

mushroom production. It is thought that low temperatures induced slow mushroom

development. Carbon dioxide levels fluctuated according to temperature and whether or

114 not mushrooms were being produced. Relative humidity was maintained at high levels,

varying between 85-95%. Figure 7.2 depicts the environmental conditions in the A. blazei tray bioreactors.

1800 35

1600 30

1400

25

1200

20 1000 CO2 Yield Temp 800 15 (ppm) & Yield (g) Yield & (ppm) 2 Temperature (C) CO

600

10

400

5 200

0 0 1234567891011121314151617181922122225227223313333533733441444454474450515253545556 Days

Figure 7.2. Environmental conditions and mushroom yield/day for A. blazei tray bioreactor

7.4.3. Agaricus bisporus

Agaricus bisporus successfully colonized and produced mushrooms in all

treatments. The effect on mushroom yield of oilseed type, rate of oilseed addition (Stage I

supplementation), and the addition of S41 (Stage II supplementation) was significant

(p<0.05). The factor interactions were mostly statistically significant (p<0.05) except for

the interaction of oilseed type x S41 addition (p>0.05). Substrates with no Stage I and

Stage II supplementation (100% millet + 0% S41) produced 4.3 kg/m2 on average,

whereas a similar substrate supplemented with 5% S41 produced 10.2 kg/m2 (statistically

significant at p<0.05). When comparing these substrates, as shown in Figure 7.3, to a

115 substrate composed of commercial grain spawn supplemented with 5% S41, the average

yield from the millet substrate colonized on-site was slightly higher although this was not

significant (p>0.05).

12 . 0 0

10 . 0 0

8.00 ) 2

6.00 Yield (kg/m Yield

4.00

2.00

0.00 0%Oil + 0%S41 0%Oil + 5%S41 Sylvan + 5%S41

Figure 7.3.Yield from two millet-grain substrates prepared on-site with one treatment receiving stage II supplementation, and compared to commercial grain spawn (Sylvan) substrate with stage II supplementation.

It was observed that a Stage I supplementation of 15% soybean with a Stage II supplementation of 5% S41 nutrient supplement at the casing stage produced 16.9 kg/m2,

which was the highest yield of mushrooms and the highest bioefficiency (205%), as

shown in Table 3. A treatment with 30% niger + 5% S41 (Stage I + II supplementation)

and another with 30% soybean + 0% S41 (Stage I) addition were high yielding as well,

116 producing 15.3 kg/m2 and 12.8 kg/m2, respectively. In almost all of the treatments, except

the 30% soybean + 5% S41treatment, mushroom yield was increased when both, Stage I

and II supplementation, were made simultaneously. Although the interaction between the

type of oilseed and S41 supplementation was not significant (p>0.05), all other

interactions such as oilseed type x oilseed rate and rate of supplementation x oilseed rate

were significant (p<0.05). Treatments with safflower produced the lowest mushroom

yield compared to the other oilseeds. Table 7.3 summarizes the data for the set of

treatments.

Table 7.3. Cultivation of Agaricus bisporus on different formulations of grain-based substrates. Treatment1 Yield2,3 Bioefficiency2,3 Average Mushroom (kg/m2) (%) Weight2,3 (g) No Oilseed 4.3 ± 0.9 a 65 ± 14 a 56.6 ± 1.2 b No Oilseed + 5% S41 10.2 ± 0.9 b,c,d,e 144 ± 14 b,c 15.1 ± 1.5 a 15% Niger + 0% S41 8.4 ± 1.7 a,b,c,d 105 ± 21 a,b 23.7 ± 7.7 a 15% Niger + 5% S41 12.2 ± 1.4 c,d,e,f 145 ± 17 b,c 19.7 ± 3.3 a 30% Niger + 0% S41 8.6 ± 1.2 a,b,c,d 92 ± 12 a,b 20.4 ± 8.7 a 30% Niger + 5% S41 15.3 ± 1.2 e,f 148 ± 17 b,c 20.0 ± 2.2 a 15% Safflower + 0% S41 5.0 ± 1.4 a,b 63 ± 18 a 25.0 ± 2.2 a 15% Safflower + 5% S41 9.0 ± 2.6 a,b,c,d 107 ± 31 a,b 20.9 ± 4.1 a 30% Safflower + 0% S41 7.3 ± 1.4 a,b,c 78 ± 14 a,b 23.0 ± 4.6 a 30% Safflower + 5% S41 8.3 ± 0.6 a,b,c,d 105 ± 7 a,b 67.0 ± 2.9 b 15% Soybean + 0% S41 10.6 ± 2.8 c,d,e 134 ± 37 a,b,c 17.8 ± 3.5 a 15% Soybean + 5% S41 16.9 ± 2.0 f 205 ± 24 c 17.5 ± 7.6 a 30% Soybean + 0% S41 12.8 ± 4.5 d,e,f 139 ± 50 a,b,c 27.4 ± 2.5 a 30% Soybean + 5% S41 8.3 ± 1.5 a,b,c,d 88 ± 16 a,b 23.1 ± 5.9 a 1Stage I supplementation is defined as the addition of oilseed to the cereal grains before spawning (15% and 30% oilseed), whereas Stage II supplementation is defined as the addition of delayed-release supplements (5% S41) to the substrate before adding the casing layer 2Treatments followed by similar letters within a column are not statistically significant (p<0.05, is significant). 3 Data presented with standard deviation.

The type of oilseed and the rate of oilseed addition had an effect on the average

mushroom size (p<0.05), whereas the effect of Stage II supplementation was not

117 significant (p>0.05). The highest average weight of mushrooms of 67 g was observed for the 30% safflower + 5% S41 (stage I and II supplementation) treatment followed by the

100% millet or no oilseed treatment + 0% S41 (56.6 g). The smallest average size mushroom of 15.1 g was observed for the no oilseed + 5% S41 treatment (Stage II supplementation).

For substrate bioefficiency, the effects of type and rate of oilseed type and S41 addition were significant (p<0.05). Oilseed type x S41 addition (Stage II supplementation) was significant (p<0.05). The highest substrate bioefficiencies were observed for the 15% soybean + 5% S41 substrate (205%), whereas the lowest was for the 15% safflower + 0% S41 (63%). Figure 7.4. depicts a treatment with A. bisporus fruiting.

.

Figure 7.4. A. bisporus fruiting on a substrate composed of grain and oilseed.

118 7.4.4. Agaricus blazei

Agaricus blazei colonized all of the substrates except for those amended with 30% soybean treatments. The effect of oilseed type and rate of oilseed addition were significant on mushroom yield (p<0.05), whereas the addition of S41 was not significant

(p>0.05). Furthermore, the interaction of oilseed type x rate of addition, oilseed type x

S41 addition, rate of oilseed addition x S41, and the three way interaction were all significant (p<0.05). Supplementing the 100% millet treatments with 5% S41 (Stage II supplementation) increased yield from 6.1 kg/m2 to 9.6 kg/m2. For all other substrates, there was a general trend of increasing mushroom yield with increasing rates of oilseeds with the exception being, as stated earlier, soybean (Table 7.4). The highest mushroom yield was obtained from the 30% niger + 0% S41 (Stage I supplementation) substrate producing 15.9 kg/m2 which was followed by the 30% safflower treatment producing

14.4 kg/m2 mushrooms. Among the lower percentage supplementation rates of oilseeds, the 15% soybean produced the highest yield. However, this was not significant (p>0.05).

When S41 was added (Stage II supplementation) to the substrate, the results differed according to the rate of the oilseed used. For the soybean treatments, supplementing with

S41 decreased the yield. However, for the 15% rate of either niger or safflower, supplementing with S41 increased average mushroom yield, which was not observed at the 30% oilseed rate of these oilseeds.

119

Table 7.4. Cultivation of A. blazei on different formulations of grain-based substrates Yield 2,3 Bioefficiency2,3 Mean Mushroom Weight2,3 Treatments1 (kg/m2) (%) (g) No oilseed 6.1 ± 2.2 b 91 ± 32 b,c 41.5 ± 6.3 b,c No oilseeed + 5% S41 9.6 ± 1.7 b,c,d,e 136 ± 8 b,c 42.0 ± 1.6 b,c 15% Niger + 0% S41 8.2 ± 0.7 b,c 102 ± 12 b,c 23.5 ± 13.9 a,b 15% Niger + 5% S41 13.2 ± 2.5 c,d,e,f 158 ± 30 b,c 26.8 ± 6.9 b 30% Niger + 0% S41 15.9 ± 0.7 c,d,e,f 170 ± 8 c 35.8 ± 1.1 b,c 30% Niger + 5% S41 13.4 ± 2.5 d,e,f 139 ± 10 b,c 39.5 ± 13.3 b,c 15% Safflower+ 0% S41 10.4 ± 0.9 b,c,d,e 130 ± 11 b,c 18.5 ± 7.8 a,b 15% Safflower + 5% S41 9.5 ± 2.6 b,c,d,e 114 ± 30 b,c 38.1 ± 7.1 b,c 30% Safflower + 0% S41 14.4 ± 1.3 e,f 154 ± 14 b,c 25.6 ± 3.3b 30% Safflower + 5% S41 8.4 ± 2.8 b,c,d 87 ± 30 b,c 41.9 ± 7.1 b,c 15% Soybean + 0% S41 13.2 ± 1.7 c,d,e,f 167 ± 21 c 31.7 ± 4.5 b,c 15% Soybean + 5% S41 6.4 ± 1.5 b 77 ± 18 a,b 51.3 ± 21.2 c 30% Soybean + 0% S41 0.0 ± 0.0 a 0 ± 0 a 0 ± 0 a 30% Soybean + 5% S41 0.0 ± 0.0 a 0 ± 0 a 0 ± 0 a 1Stage I supplementation is defined as the addition of oilseed to the cereal grains before spawning (15% and 30% oilseed), whereas Stage II supplementation is defined as the addition of delayed-release supplements (5% S41) to the substrate before adding the casing layer 2Treatments followed by similar letters within a column are not statistically significant (p<0.05, is significant). 3 Data presented with standard deviation.

All three main effects (oilseed type, oilseed rate, and S41 supplementation) were

significant (p<0.05). Also, all the interactions among the effects were statistically

significant. However, it was observed that the greater mushroom yields correlated with a

larger number of mushrooms. This was not the case for the 15% safflower + 0% S41

substrate (Stage I supplementation), which produced the highest number of mushrooms,

but was a lower yield compared to the 30% safflower + 5% S41 (Stage I and II

supplementation) treatment. However, the highest mean mushroom weight was observed

for the 15% soybean + 0% S41 (Stage I supplementation) substrate. Figure 7.5 depicts a

treatment with A. blazei fruiting.

120

Figure 7.5. A. blazei mushroom fruiting on a grain/oilseed substrate

7.5. Discussion

Non-composted substrates for the production of Agaric mushrooms is a research focus that warrants further attention. Both A. bisporus and A. blazei can be grown successfully on non-composted substrates. Furthermore, both mushrooms are well known to require a non-sterile casing for fruiting (Stamets, 2000b). For A. bisporus, several studies showed that the addition of activated carbon to sterilized casing restored mushroom production (Long and Jacobs, 1974; Verbeke and Overstyns, 1991), whereas this is the first report of such a study for A. blazei. In our study, a non-composted substrate composed of a mixture of grains and oilseeds was tested. It is hypothesized that the grains provide the mushroom fungus with the carbohydrates and water through the

121 gelatinization of starch, whereas the oilseeds satisfy the lipid and protein requirement.

Interestingly, Dahlberg (2004) recently provided evidence suggesting that carbon limits

the production of mushrooms, because an increase in yield was directly related to an

increase in the quantity of carbohydrate added to the compost. This raises the question of

whether it is the carbon and not the nitrogen component, as it has long been viewed, in

delayed-release supplements that stimulates yield.

A comparison of the different substrate treatments provided evidence that the two mushroom species have different preferences. Overall, the yield of A. bisporus on the

millet-grain substrate was comparable to that observed for commercial rye grain spawn.

In our trials using substrates composed of grain and oilseeds with A. bisporus, there was a

tendency for increased mushroom yield with the addition of soybean relative to safflower

and, to some extent, niger. Furthermore, a general trend of increasing yield for A.

bisporus with the use of S41 in Stage II supplementation was observed. However, this

was not the case for the treatment containing 30% soybean + 5% S41. Furthermore,

mushrooms produced from the 30% soybean treatments formed mostly in clusters and

were highly deformed, which suggests toxicity at this higher rate. Overall, there is

sufficient evidence to conclude that adding oilseeds to the substrate prior to the growth of

A. bisporus, Stage I supplementation, as well as the addition of a delayed-release nutrient

supplement at the casing stage, (“Stage II supplementation”), increases mushroom yield.

Substrate bio-efficiencies were also high when compared to traditional substrates.

Composted substrates with bio-efficiencies ranging between 70-90% are considered good

(Schisler, 1982). The observed bio-efficiencies for the grain and oilseed substrates were

122 considerably higher than composted substrates due to the fact that the nutrients are supplied to the mushroom fungus in greater concentrations.

Mean mushroom weight was also evaluated in this study and, generally, low yielding substrates produced larger mushrooms. However, there were some exceptions, especially considering the high mean mushroom weight for the 30% safflower + 5% S41 treatment. This is significant because mushroom quality is an important determinant of market value. Larger mushrooms command higher prices than smaller ones allowing mushroom producers to profit even further. More work is needed to better understand the relationship between grain/oilseed substrate formulations and mushroom size, quality, shelf-life, and ultimately profit.

In A. blazei trials, there was generally increased mushroom yield in treatments with niger and safflower relative to substrates with millet grain and no oilseeds, whereas reduced yields were obtained from treatments with soybean (Stage I supplementation).

In fact, complete inhibition was observed at the 30% soybean rate. Interestingly, S41 increased yield for millet + no oilseed and substrates containing 15% niger or safflower.

This was not the case for the 15% soybean + 5% S41 or the higher 30% niger and safflower treatments supplemented with S41. Bioefficiencies values of A. blazei were reported to be between 50-75% on composted substrates (Stamets, 2000b). As in the case of A. bisporus, the bio-efficiencies of the treatment substrates were far greater than those reported for composted substrates. Mean mushroom weight varied as well. However, in contrast to A. bisporus, A. blazei tends to be sold in a dehydrated form or as capsules so total yield may be far more important than mean mushroom weight.

Cultivating Agaric mushrooms on non-composted substrates eliminates many of the

123 environmental issues now confronting the commercial mushroom industry. Composting,

although an economical method of producing a substrate for Agaric mushrooms, is labor intensive, time consuming, and necessitates the use of heavy equipment for handling the

bulky material. Most importantly, composting within the mushroom production process

generates unpleasant odors, nutrient-rich run-off, and a waste stream (spent substrate)

that also emits odors and is difficult to eliminate (Heinemann, et al., 2003; Heinemann et al., 2004). Substrates composed of cereal grains and oilseeds have the potential of serving as substrates for commercial mushroom production. On average, a grain-base substrate is less bulky than traditional compost, because the seed is highly concentrated with nutrients. Furthermore, the substrate preparation process can be entirely automated, thereby reducing the cost of labor and heavy machinery use. Finally, as for the waste stream, spent grain and oilseed substrates, might be marketed as animal feed.

7.6. Conclusions

Agaricus bisporus and A. blazei mushrooms were successfully produced on

sterilized substrates consisting of millet grains and different types of oilseeds. The

addition of S41, a commercially available delayed-release supplement for compost, to the

substrate prior to the addition of casing (Stage II supplementation) increased the yield of

A. bisporus in almost all substrates but not for A. blazei. Also, A. blazei failed to colonize

substrates containing 30% soybean, whereas A. bisporus colonized these substrates, but

developed malformed mushrooms. Overall, mushroom yield was influenced by the type

and rate of oilseed mixed with millet grain prior to colonization by the mushroom fungus and by the further addition of S41 after colonization of the substrate. The highest yield of mushrooms for a 2-2.5 cm depth of grain/ oilseed substrate for A. bisporus was 16.9

124 kg/m2, whereas the commercial yield from conventional composted substrates (~0.2 m depth) is around 28.8 kg/m2. The highest yield obtained for a grain/oilseed substrate with

A. blazei was 15.9 kg/m2, which compares favorably with reported yields in the range of

17 kg/m2 on composted substrates

Acknowledgments

The authors would like to acknowledge the assistance of Tom Rhodes, manager of the Penn State mushroom research facilities. Furthermore, the authors would like to thank Dr Eileen Wheeler for supplying the carbon dioxide measuring device. Funding was provided by the College of Agricultural Sciences and the Department of Agricultural and Biological Engineering, The Pennsylvania State University.

125 Chapter 8

Pre-incubating Non-composted Grain Substrates with the Thermophilic Fungus Scytalidium thermophilum for Mushroom (Agaricus bisporus) Production

8.1. Abstract

Grain-based substrates for Agaricus bisporus were tested as alternatives to the

environmentally problematic compost-based substrate. Scytalidium thermophilum, the dominant thermophilic fungal species found in compost-based mushroom substrates, was used to pre-treat grain-based substrates in an effort to reduce spawn-run duration, increase mushroom production, and improve substrate bioefficiency. Three sterilized substrates composed primarily of grains (rye, millet, and hulled oats) with a mix of cracked roasted soybean and other lesser ingredients were used as the basal substrate and were either inoculated with S. thermophilum for various days ranging from 0-20 days at

46oC or not. Results indicated that the incubation period had a significant effect on

mushroom yield and bioefficiency (p<0.05). Yield decreased significantly beyond the 10

day incubation period when compared to the control treatments. As a result of the

preliminary treatments, shorter incubation durations (0, 2, 4, and 6 days) at 46oC were

recommended. Spawn-run durations for substrates inoculated with S. thermophilum were

shorter (22-23 days) compared to control treatments (44-50 days) and the interaction of

grain type and incubation duration was the only significant factor (p<0.05). Overall, S.

thermophilum can effectively reduce spawn run duration and improve mushroom yield

for oat-based substrates.

Keywords Thermophilic fermentation, solid-state fermentation, mushroom production.

126 8.2. Introduction

Production of the widely cultivated button mushroom (Agaricus bisporus) on

partially composted substrates is a mainstay in industry. Although there are several

variations in the substrate preparation process such as the long method, INRA method,

and the Anglo Dutch method (Rai, 2004), the most widely used process in Pennsylvania

was developed by Sinden and Hauser (1950) and called the “short method”. The short

method consists of two phases, whereby in the first phase (Phase I) raw ingredients

(horse manure, chicken manure, hay, straw, gypsum and others) are mixed wetted and

placed in rows outdoors for uncontrolled self heating, and takes 7 -14 days for

completion. Next, the materials are transferred to temperature controlled

chambers/tunnels in which the partially composted material is pasteurized to inactivate

mushroom pathogens, and maintained at 45-50 °C for about 5-7 days. During this phase,

thermophilic microorganisms grow rapidly and transform the substrate into a mushroom-

specific substrate i.e. free of pathogens of A. bisporus. This concludes the second phase

(Phase II) of the substrate preparation step after which the partially composted material is

termed “mushroom substrate” and is ready for spawning (inoculation with mushroom

grain spawn). In summary, the substrate preparation process confers selectivity of the substrate A. bisporus growth while providing an adequate balance of nutrients and water

to sustain mushroom production. During the temperature control period of Phase II, the

dominant thermophilic species is Scytalidium thermophilum a fungus (Straatsma et al.,

1994).

Cooney and Emerson (1964) state that the optimal temperature for S. thermophilum growth is around 40 °C, whereas the maximum temperature for growth is

127 58 °C. Fergus and Amelung (1971) and Satyanarayana and Johri (1984) report that

conidiating colonies of S. thermophilum are inactivated at 68 °C. S. thermophilum was

reported to grow better under anaerobic or micro-aerobic conditions than under aerobic

conditions at elevated temperatures (Henssen,1957; Maheshwari et al., 2000). The effect

of S. thermophilum and the thermophilic biomass found in compost-based substrates on

A. bisporus was discussed in several studies.

Inoculating the substrate with S. thermophilum increased mushroom yield two-

fold (Straatsma et al., 1994), and other studies have shown that the selectivity of the substrate is attributed to the thermophilic microflora which dominates the substrate at spawning (Ross and Harris,1983; Wiegant et al., 1992). Ross and Harris (1983) showed that heat and antimicrobial chemicals destroy the selectivity of partially composted substrate by killing the thermophilic microflora. Therefore selectivity of the compost is related to a living biomass. They go on to explain that the reduction in temperature of

mushroom substrate after Phase II produces “static” conditions (dormant) and not

“killing” conditions in which the thermophilic microflora remain viable ensuring the

selectivity of the substrate for A. bisporus. Wiegant et al., (1992) showed that a significant lag in biomass formation of the mushroom fungus was due to low carbon dioxide concentrations in the sterile compost compared to non-sterile compost. Wiegant et al. (1992) also showed that sterilized compost failed to give rise to mushroom mycelium growth. However, the addition of S. themophilum to the sterilized material

either pre-incubated or at the time of spawning led to the same mycelium extension rate

with the latter characterized by a lag phase. Furthermore, carbon dioxide was shown to be

an important factor on extension rates, whereby 0.5% concentrations sustained an

128 increase in extension rates and above which extension rates leveled off. Increase in

mushroom mycelium extension rates was also reported for CO2 concentrations of up to

1% (San Antonio and Thomas, 1972). The explanation behind the stimulating effect of

CO2 was described as an ecological effect, whereby the mushroom fungus senses the

growth of other fungi and tries to colonize the substrate as rapidly as possible to ensure

its survival (Wiegant et al., 1992). However, the more probable reason for CO2 stimulation is its role in anaplerotic reactions in which carbon fixation occurs for the formation of intermediate compounds in the TCA cycle and in the formation of purines pyrimidines and fatty acids (Maheshwari et al., 2000; Carlile et al., 2001). The major contribution of carbon dioxide at cultivation temperatures of 24 °C was shown to be mainly due to the growth of the thermophilic fungi and less to the prokaryotic microflora

(Wiegant et al., 1992). Wiegan et al., (1992) state that although high carbon dioxide concentrations stimulated extension rates, the growth promoting effect of S. thermophilum either by secreting stimulating compounds or vital nutrients cannot be

ruled out. S. thermophilum produces large quantities of cellulases and xylanases that

could aid in the substrate pre-conditioning (Zanoelo et al., 2004a; Zanoelo et al., 2004b).

Although widely used, the traditional substrate preparation process has come

under a lot of scrutiny due to the production of odors and nutrient-rich runoff and

leachate. Furthermore, the amount of spent mushroom substrate generated in some areas is too large for land disposal. Non-composted substrates have been shown to be adequate

replacements for the environmentally problematic composted substrate.

Growing the mushroom fungus on grains is mainstay in the spawn (the vehicle

used to inoculate traditional composted substrates) industry. A previous study by Bilay

129 (2000) pre-incubated sterilized grain with H. insolens (S. thermophilum) and then inoculated the colonized grains with A. bisporus to form what was called as

“experimental grain spawn” (EGS). Bilay (2000) states that the advantage of using EGS

is in reducing the problem of mold contamination. San Antonio (1971) showed that A.

bisporus mushroom was possible on a substrate composed primarily of rye grains.

Bechara et al. (2006a) showed that adequate mushroom production can be achieved by

using a 2 cm layer of sterilized substrate composed of millet grain and various oilseeds

and observed yields were as high as 16.9 kg/m2. The colonization phase duration of the

mushroom fungus mycelium took 32 days, which is far greater than the typical

colonization phase of 14 days reported for composted substrates.

The objective of this is study was to determine whether S. thermophilum could be

used as a conditioner for non-composted grain-based substrates that could effectively

decrease spawn-run duration, increase mushroom yield and substrate bioefficiency.

8.3. Methods and Materials

8.3.1. Fungal Cultures Substrate Materials

Rye grain spawn cultures of Scytalidium thermophilum (DC 295) and Agaricus bisporus (MC 459) were obtained from the Mushroom Spawn lab (Penn State

University). S. thermophilum was stored at 37°C, whereas A. bisporus was kept at 5 °C.

Next, the grain substrates were composed of a grain portion (hulled oats, millet, and rye)

mixed with 40 g cracked roasted soybean (Agway, Pleasant Gap, PA) + 10 g CaCO3 and

10 g CaSO4 (Fisher Scientific, Hampton, NH) + 15 g DAC vitamin/mineral supplement

(DAC, Dover, OH) and either 2000 ml or 500 ml of perlite, a water-holding material. Rye

grains were supplied from the Mushroom Spawn Lab, whereas hulled oats and millet

130 were purchased from a local feed store (Agway, Pleasant Gap, PA). Perlite (Therm-o-

Rock, New Eagle, PA) was also added to the mushroom production trays.

8.3.2. Preparation of Grain Substrates

Grains and the appropriate amount of water to achieve a final moisture content of

0.6 (kg/kg wet basis) were added to polypropylene (L = 0.37 m x W = 0.25 m x D = 0.14 m) containers. The filled containers were covered with a double layer of aluminum foil, autoclaved for 45 minutes, and allowed to cool to room temperature. Following, the grain substrates components were mixed and added to Unicorn® (Unicorn Imp. & Mfg. Corp.,

Commerce TX) type 10 autoclavable polypropylene bags with filter patches containing

2000 ml or 500 ml perlite. Perlite was added to the bags to absorb the extra condensate evolved from the thermophilic biomass.

The filled bags were then autoclaved for 3 hrs, and allowed to cool in a laminar flow hood. Once cooled, the bags were aseptically inoculated with 20 g of rye grain spawn (S. thermophiluim and/or A. bisporus). Substrates inoculated with S. thermophilum were incubated at 46 °C for the treatment incubation duration and then

inoculated with A. bisporus after cooling down to room temperature. Substrates

inoculated with A. bisporus, or a co-culture of A. bisporus and S. thermophilum (0 day

treatment) were incubated at 22 °C. Once fully colonized, the mushroom substrates were

transferred to mushroom production trays.

8.3.3. Preparation of Mushroom Production Containers

Autoclaved (121 °C for 60 min) polypropylene containers (0.3 m, 0.16 m, 0.09 m)

were used as mushroom production trays. Perlite was wetted and each mushroom tray

received 2000 ml of perlite. The containers were then autoclaved for 45 min (121°C and

131 103.4 kPa) and then allowed to cool. Mushroom casing (lime neutralized peat moss) was

obtained in dried form from the Mushroom Research Center (The Pennsylvania State

University). The casing was wetted (7000 ml casing + 5000 ml hot tap water) and then supplemented with 10% activated carbon v/v (Fisher Scientific, Hampton, NH). The casing was autoclaved (121°C and 103.4 kPa) for 2 hrs.

The colonized grain substrates were added to the mushroom production trays and treatments receiving delayed-release supplements were amended with 40 g of Promycel

Target® (SpawnMate, Watsonville, CA). The sterilized casing (limed peat moss with activated carbon) was then added over the substrate (1200 g/ tray). Mushroom production

was carried out in a mushroom production chamber.

8.3.4. Environmental Conditions in Mushroom Production Chamber

Filled and cased mushroom production trays were placed in a tray bioreactor

(2.54 m x 1.32 m) at the Agricultural and Biological Engineering Department.

Temperature was maintained using a commercial air conditioning unit. Three indoor

greenhouses (Midwest Quality Qlove Inc, Chillicothe, MO) were used to store the

mushroom trays in the tray reactor, and they consisted of metal structures with four

shelves that could hold up to five trays. The structures were enclosed within a

polyethylene cover. Slits measuring 9 cm were made in the polyethylene covers to

improve ventilation. Two slits were made on the top of the chambers and the rest were

made on the front side. The temperature was maintained at 22 oC. The temperature was

then dropped to 16 oC and maintained until mushroom harvesting was completed. \

132 8.3.5. Experimental Set-up and Analysis of Data

Two sets of experiments were undertaken. The first set of treatments were made

based on a randomized complete block design (block = time) and tested two factors:

incubation duration at 46 oC for 0, 10, and 20 days, and type of grain used (rye, millet,

and oat). The treatments were compared to a control group with no S. thermophilum

addition. Table 8.1 provides a summary of the first experimental setup.

Table 8.1. Treatments with different high temperature incubation periods with and without supplementation with Promycel Target® Incubation Promycel Second Stage Treatment1,2 First Stage Inoculation at 46oC Target Inoculation (days) (g) Ctrl A. bisporus 0 - 0 0 day A. bisporus + S. 0 - 0 thermophilum 10 day S. thermophilum 10 A. bisporus 0 20 day S. thermophilum 20 A. bisporus 0 1 Each bag contained 2000 ml perlite and 835g of substrate. 2 For each incubation duration the three types of grains (rye, millet, and hulled oats) were used.

The second set of treatments was made using a completely randomized design with incubation duration (0, 2, 4, and 6 days) and grain type (rye, millet, and hulled oat) as factors. All treatments in the second experiment were supplemented with Promycel

Target®. Table 8.2 summarizes individual treatments performed in the second experiment. For both set of experiments, treatments were replicated thrice and mushrooms were harvested for a period of 26 days.

133

Table 8.2. Treatments with shorter high temperature incubation periods all supplemented with delayed-release supplement Promycel Target® Incubation Promycel Second Stage Treatment1,2 First Stage Inoculation at 46oC Target Inoculation (days) (g) Ctrl A. bisporus 0 - 40 A. bisporus + S. 0 day 0 - 40 thermophilum 2 day S. thermophilum 10 A. bisporus 40 4 day S. thermophilum 10 A. bisporus 40 6 day S. thermophilum 20 A. bisporus 40 1 Each bag contained 500 ml of perlite with 835g of substrate. 2 For each incubation duration the three types of grains (rye, millet, and oats) were used.

The response variables recorded and calculated were mushroom yield, spawn-run duration, average mushroom size, and substrate bioefficiency. Spawn-run duration is defined as the number of days it takes to fully colonize the substrate (grains are fully covered with biomass) with mycelium and have it ready for transfer to mushroom

production trays. Average mushroom size was calculated by dividing the total mushroom yield (fresh weight) by the number of mushrooms picked, whereas substrate bioefficiency

was calculated by dividing the total mushroom yield (fresh weight) by the oven-dry

weight of the substrate components. Perlite was not counted as part of the substrate and

was not included in the bioefficiency calculations. The response variables were

statistically analyzed using the General Linear Model (α = 0.05) with MINITAB statistical software (release 13.1). All pairwise comparisons of treatments were made using the Tukey method.

134 8.3.6. Oxygen Uptake Rate Evaluation

Oxygen uptake rates (OUR) of S. thermophilum and A. bisporus at 16, 24, and

32oC were measured using the Oxitop® system (WTW, Giessen, Germany). The system

is composed of a 1.14 L glass bottle that is partially filled with a biomass sample and a

bottle sealing cap composed of a receptacle for a carbon dioxide absorbent (typically

NaOH) and a pressure sensor head that transmits the enclosed bottle pressure reading to a

controller. The OUR can be calculated based on the pressure drop in the apparatus. The

pressure drop is induced by the oxygen consumption of the living biomass. The Oxitop®

system has been successfully used in several studies for the measurment of compost and

compost bulking material OURs (Sadaka et al., 2004; Ahn et al., 2005).

For this study, a 50 g sample of rye grain pre-colonized with either A. bisporus or

S. thermophilum were added to the Oxitop® system. The bottles and receptacales were

sterilized with the exception of the pressure sensor head. Next, 2 g of concentrated NaOH

pellets were added to the CO2 absorbent receptacle and the bottles cap was sealed with

the pressure heard sensor. The treatment duration was 48 hrs. Dry matter content was

measured by taking a 10 g sample of the grain spawn and placing them in an oven at

105oC for 24 hrs. Pressure drop data were converted into OUR as shown in equation 1.

⎛ hPa⎞ ⎛ Pa ⎞ 3 slope⎜ ⎟ ×100⎜ ⎟ ×Vgas()m × 32()g/moleO2 ×1000()mg/g ×1440() min day ⎛ mgO ⎞ ⎝ min⎠ ⎝ hPa⎠ OUR⎜ 2 ⎟ = Eq. 8.1 ⎝ gDMday⎠ ⎛ J ⎞ 8.314⎜ ⎟ × TK()×Wg()× (1 − MC) ⎝ moleK ⎠ where slope = slope of the pressure drop curve

Vgas = volume of gas occupying the Oxitop vessel- assumed total volume of flask.

T = temperature

135 W = weight of sample

MC = moisture content wet basis

the remaining numbers in Equation 8.1 are conversion factors.

8.4. Results

8.4.1. Preliminary Incubation experiments with S. thermophilum

For the first trials with S. thermophilum addition, the only significant effect on

mushroom yield was incubation duration (p<0.05). Yield for the different grain-based substrates was not significantly different (p>0.05), and neither was the interaction effect grain x incubation duration (p>0.05). Overall, yield for the 0 and 10 day were not significantly different from the control group, as indicated in Table 8.3. However, yield was significantly lower for the 20 day incubation duration, and some replications of the

20 day incubation duration failed to grow. The grain spawn when added to the bags was inactive for weeks, and mushrooms were not produced when treatments were transferred to trays. An observed decrease in spawn-run duration was detected in treatments

inoculated with S. thermophilum compared to the control. Furthermore, average

mushroom yield was lower for the 10 day incubation duration as compared to the control

and 0 day incubation duration, and so the incubation duration was reduced. The highest

yield was for the 0 day rye treatment producing 19.4 kg/m2 with a corresponding BE =

249.3%. Table 8.3 provides a summary of results for the 0-10- and 20 day incubation

durations. For average mushroom size, none of the factors or interaction effect were significant (p>0.05)

136

Table 8.3. Preliminary treatments for incubation of grain-based substrate with S. thermophilum Average Yield Biological Efficiency Mushroom Size Treatment1,2,3 (kg/m2) (%) (g) Ctrl- Rye 16.6 ± 8.1a 213.4 ± 104.1a 16.2 ± 2.4a Ctrl- Millet 15.2 ± 2.5a,b,c 195.0 ± 32.5a,b,c 20.9 ± 4.9a Ctrl- Oats 15.4 ± 1.5a,b,c 197.1 ±19.1a,b,c 18.7 ± 1.7a 0 day- Rye 19.4 ± 6.3a 249.3 ± 80.7 a 21.1 ± 5.4a 0 day- Millet 16.2 ± 6.5a,b 207.8 ± 83.4a,b 18.6 ± 1.0a 0 day- Oat 18.6 ± 2.9a 238.3 ±37.4a 17.7 ± 1.0a 10 day- Rye 9.3 ± 4.1a,b,c,d 118.7 ± 52.5a,b,c,d 15.5 ± 0.6a 10 day- Millet 10.2 ± 4.9a,b,c,d 130.8 ± 63.9a,b,c,d 17.6 ± 3.7a 10 day- Oat 8.5 ± 4.5a,b,c,d 109.0 ± 57.7a,b,c,d 15.3 ± 1.7a 20 day- Rye4 2.8 ± 2.9c,d 35.9 ± 37.2c,d 15.8 ± 16.4a 20 day- Millet4 3.4 ± 5.4c,d 43.6 ± 69.2c,d 32.4 ± 17.4a 20 day- Oat4 1.5 ± 1.5d 19.2 ± 19.2d 4.1 ± 7.0a 1 Data presented with standard deviations 2 Treatments followed by similar letters are not significantly different (p<0.05, is significant) 3Observed data is for 26 days of harvesting. 4Treatments with large standard deviation because one replicate failed to yield any mushrooms.

8.4.2. Treatments with Shorter Incubation Durations with S. thermophilum

8.4.2.1. Spawn-run Duration

Pre-incubating S. thermophilum and then inoculating with A. bisporus or

simultaneous inoculation with S. thermophilum and A. bisporus reduced the spawn-run

duration from 44-50 days for the control treatments to 22-23 days for the 6 day

incubation duration. The longer the thermophilic incubation period, the shorter the

spawn-run duration. The shortest spawn-run duration was observed for treatments with

the 6 day incubation period with S. thermophilum, whereas the longest spawn-run

duration was observed for the control treatment with no S. thermophilum addition. Table

4 summarizes the spawn run duration for all the treatments.

137

Table 8.4. Colonization phase duration as influenced by the addition and pre-colonization of S. thermophilum in grain-based substrates Spawn-run Duration Total Duration3 Treatment1,2 (days) (days) Ctrl- Rye 49 ± 2a 49 Ctrl- Millet 44 ± 8a,b 44 Ctrl- Oats 50 ± 4a 50 0 day- Rye 48 ± 0a 48 0 day- Millet 28 ± 2b,c,d 28 0 day- Oat 39 ± 8a,b,c,d 39 2 day- Rye 30 ± 5b,c,d 32 2 day- Millet 38 ± 9a,b,c,d 40 2 day- Oat 35 ± 12a,b,c,d 37 4 day- Rye 27 ± 4b,c,d 31 4 day- Millet 24 ± 2d 28 4 day- Oat 26 ± 5c,d 30 6 day- Rye 22 ± 2d 28 6 day- Millet 23 ± 2d 29 6 day- Oat 22 ± 2d 28 1 Data presented with mean ± standard deviation 2 Treatments followed by similar letters are not significantly different (p>0.05) 3 Total duration includes the days of incubation at 46oC and spawn-run

8.4.2.2. Yield, Bioefficiency, and Average Mushroom Size

The interaction effect (grain type x incubation duration) and incubation duration

were the only significant terms for mushroom yield (p<0.05). Grain type as main effect

was not significant (p>0.05). Yield for the oat-based substrate resembled a bell shaped

curve that peaked at the 2-day incubation duration. The rye substrate did not have a clear peak in yield but the highest yield was for the 0 day incubation duration. Finally, for the

millet substrate the highest yield was observed for the control group which was then

lower for all other incubation durations. The highest mushroom yield was observed for

the millet control treatment, producing 21.3 kg/m2 BE = 273% as shown in Table 5.

138 Grain type and incubation duration did not significantly influence average mushroom size (p>0.05). However, the interaction grain type x incubation duration was significant (p<0.05) on average mushroom size. Average mushroom size increased with incubation period for oats and rye. In contrast, it decreased for millet. The highest observed average mushroom size was for the millet control treatment (53.5 g) and the lowest was observed for the 6 day oat treatment (21 g).

Table 8.5. Incubation duration and grain type and their influence on mushroom yield, substrate bioefficiency, and average mushroom size Yield Bioefficiency4 Average Mushroom Treatment1,2,3 (kg/m2) (%) Size (g) Ctrl- Rye 13.9 ± 4.9 b 179.4 ± 62.8b 27.7 ± 12.5a,b Ctrl- Millet 21.3 ± 0.6 a 273.4 ± 8.3a 53.5 ± 8.5a,b Ctrl- Oat 11.7 ± 3.0 b 150.5 ± 38.6b 25.0 ± 15.5b 0 day- Rye 15.6 ± 2.4a,b 200.4 ± 31.0a,b 28.7 ± 3.8a,b 0 day- Millet 13.1 ± 2.3b 167.9 ± 30.1b 27.7 ± 10.2a,b 0 day- Oat 15.6 ± 1.1a,b 200.1 ± 14.4a,b 44.3 ± 3.1a,b 2 day- Rye 15.2 ± 1.6 a,b 194.8 ± 20.2a,b 37.7 ± 10.9a,b 2 day- Millet 15.9 ± 1.1 a,b 204.5 ± 14.4a,b 36.0 ± 6.0a,b 2 day- Oat 18.2 ± 0.2 a,b 233.4 ± 3.2a,b 59.7 ± 6.5a 4 day- Rye 13.1 ± 1.2 b 168.0 ± 15.7b 31.3 ± 9.7a,b 4 day- Millet 14.2 ± 2.6 a,b 182.6 ± 33.2a,b 45 ± 25.2a,b 4 day- Oat 15.8 ± 4.6 a,b 201.9 ± 58.7a,b 35.7 ± 17.6a,b 6 day- Rye 13.5 ± 1.6 b 173.4 ± 20.2b 32 ± 4.6a,b 6 day- Millet 15.0 ± 3.3 b 192.4 ± 41.7b 42.7 ± 3.8a,b 6 day- Oat 11.2 ± 2.0 b 143.6 ± 25.9b 21 ± 8.7b 1 Data presented with standard deviations 2 Treatments followed by similar letters are not significantly different (p<0.05, is significant) 3Observed data is for 26 days of harvesting.

8.4.3. Oxygen Uptake Rate Evaluation

The OUR of A. bisporus and S. thermophilum were measured and the observed trend was anticipated, whereby a steady increase in OUR was observed for both fungal species until 32 oC. At 32 oC the OUR of S. thermophilum continued to increase, whereas

139 that of A. bisporus decreased because it is closer to the upper temperature limit of growth.

Figure 1 depicts the general trend of OUR for both fungal species.

25

20 )

15 Scytalidium A. bisporus 10 OUR (mgOUR O2.DM-1.day-1

5

0 16 24 32 Temperature (C)

Figure 8.1. OUR measurements for A. bisporus and S. thermophilum at 16, 24, and 32oC.

8.5. Discussion

Pre-incubating and co-cultivating S. thermophilum in non-composted grain-based

substrate for A. bisporus mushroom production decreased spawn-run duration for all

grain-based substrates, whereas yield and substrate bioefficiency improved for some

grain-based substrates. This is the first report of testing three different grains (rye, millet,

and oat) for mushroom production, and the addition of S. thermophilum as a pre-

conditioning agent to such substrates. Earlier, work showed that grain-based substrates

(commercial grain rye and millet grain spawn, or millet substrate supplemented with

oilseeds) supported high mushroom yields (16.9 kg/m2) but comparatively lower than

composted substrate yields (Bechara et al., 2005b; Bechara et al., 2006b). However,

140 substrate bioefficiencies for the grain-based substrates were much higher than for

compost-based substrates. In effect, adopting a grain-based mushroom production system could effectively reduce the mushroom production process by at least 21-30 days with the elimination of composting as a substrate preparation step. However, spawn-run durations for the grain-based substrates as shown in this study, are much higher (44-50 days) compared to composted substrate (14 days). Therefore, pre-incubating the grain-based substrate with S. thermophilum is an adequate solution to reduce spawn-run duration, and

ultimately reducing the total mushroom production process for grain-based substrates.

Other methods of reducing spawn-run duration are increasing inoculum amount/unit mass

and thu increasing the number of inoculation points. An efficient way of increasing

inoculation points is using smaller particles or carriers such as millet grain spawn or

casing inoculum. Casing inoculum is typically a vermiculite-based material covered with

A. bisporus mycelium that is added to mushroom casing and not mushroom substrate to

reduce spawn-run duration in traditional compost-based mushroom production (Mamiro,

2007). In addition, there was no difference in yield between the three different grains

used. Hence, a mushroom producer can use the cheapest source of grains for mushroom

production. In this study, oat was the cheapest grain among the three tested. An important

factor observed to influence growth of A. bisporus in substrate pre-incubated with S. thermophilum was the addition of perlite to the substrate itself. A. bisporus failed to

colonize the pre-incubated substrates when perlite was absent because condensate would

accumulate at the bottom of the bag, leading to a loss of structure and increase in

substrate stickiness. This was not apparent for control treatments lacking S. thermophilum, but was found in the 0, 2, 4, 6, 10, and 20 day treatments.

141 S. thermophilum, although a thermophile, appears to grow adequately at both

24oC and 16oC indicating that fungus actually is still viable and the carbon dioxide output

from the thermophilic biomass at both temperatures is a contributing factor for the

reduction of spawn run durations as many other studies have indicated. Hence, adding S. thermophilum to the sterilized grain-based substrate without the high temperature incubation period may prove to be just as effective as the 2-4 and 6 day high temperature incubation durations. More work is needed to optimize substrate composition to further increase yield and bioefficiency.

8.6. Conclusions

Mushroom yield, substrate bioefficiency, and spawn-run duration in non- composted grain-based substrates were enhanced to varying degrees with the addition of

S. themophilum. Hulled Oats, millet, and rye grain substrates were all shown to support

high mushroom yields (17.9- 21.3- 19.4 kg/m2, respectively), yet still lower than the

norm from commercial operations using compost-based substrates which on average

produce 30 kg/m2. One important observation is the decrease in spawn-run durations for

treatments with S. thermophilum, compared to controls without S. thermophilum which

ultimately shortens the production cycle. The effect of S. thermophilum in these different

grain substrates was dependent on the type of grain used. Oat-based substrates benefited

the most from the addition of S. thermophilum i.e. mushroom yield and bioefficiency

increased, whereas rye and millet-based substrates did not. In summary, the addition of S.

142 thermophilum to grain-based substrates can be used to shorten spawn- run durations and improve mushroom yield for some grain-based substrates.

143 Chapter 9

Effect of Substrate Moisture Content on Agaricus bisporus Oxygen Consumption Rate, Hyphal Extension Rate, Mushroom Yield, and Bioefficiency

9.1. Abstract

Production of Agaricus bisporus mushroom was tested on rye-grain based

substrates as an alternative to the environmentally problematic composted substrates. Rye

grain substrate moisture content was varied (45-50-55-60-65- and 70%) and the effect of

moisture content on A. bisporus oxygen uptake rate (OUR), hyphal extension rate,

mushroom production, and bioefficiency was tested. Furthermore, the effect of

measurement frequency on OUR was tested. Overall, A. bisporus OUR and hyphal

extension rates increased with increasing moisture levels. The highest OUR of 4.88 mg

O2/g VS day was observed for the 65% moisture level and the lowest (1.96 mg O2/g VS day) for the 45% moisture level. Frequency of measurement had an effect on OUR with the higher frequency lowering the OUR compared to the lower measuring frequency. It is recommended that continuous monitoring of fungal OUR as opposed to manometric methods, be used for these types of tests to eliminate this problem. Moisture level had an effect on mushroom yield and substrate bioefficiency but not in all experiments.

However, on average moisture levels ranging from 50-65% are recommended. Both the

45% and 70% moisture level were problematic because of increased probability of failure

(no growth) and at the 70% moisture level, substrate stickiness.

144 9.2. Introduction

Commercial Agaricus bisporus mushroom production is achieved using a basal medium composed of organic matter that is composted and amended with a protein-rich delayed-release supplement (Bechara et al., 2006a). Average mushroom yield for the

USA is 30 kg/m2 (USDA 2005-2006). Although widely adopted in the mushroom

industry, the use of composted substrates is a potential nuisance for mushroom producers

because of odor production and nutrient-rich run-off from substrate preparation sites.

Another concern for mushroom producers is the generation of a large organic waste

stream (spent mushroom compost) that has not been biologically-stabilized, i.e. resumes

producing odors, and requires permits for land-application (Heinemann et al., 2003;

Heinemann, et al., 2004). Thus, many efforts and studies are being made to circumvent

the need for compost-based substrate by replacing them with non-composted substrates

for Agaricus mushroom production.

Recent studies on non-composted substrates have focused on substrate

formulations composed of lignocellulosic waste products with other lesser ingredients, and grain-based substrates (Sanchez and Royse, 2001; Mamiro et al., 2007; Bechara et al., 2006ab; Sanchez et al., 2007). The production of Agaricus mushrooms on both

composted and non-composted substrates is a solid-state fermentation process (SSF). SSF is defined as the growth of microorganisms–fungal organisms in the case of mushroom production–on a solid-matrix in the absence of free-flowing water (Pandey, 2000).

Fungal growth in SSF is influenced by various environmental and process parameters such as temperature, pH, substrate moisture content, and aeration (Raimbault, 1998).

Typically, the solid-matrix is both the source of nutrients and water necessary for the

145 growth of microorganisms. Therefore, an important factor that influences the growth of

microorganisms is the moisture content within the solid matrix. Substrate porosity, the

volume fraction of pores in a solid matrix, is an important variable that affects the growth

microorganism. Pores can be filled with either air (air-filled porosity) or liquid, and if

filled with liquid, then oxygen diffusion is hindered. Air filled porosity is critical since

most fungi, including A. bisporus, are aerobic microorganisms, i.e. have oxygen as the

terminal electron acceptor. Thus, the availability of oxygen is important and its

limitations will lead to lower or no growth. Aside from oxygen within the air-filled pore

space, carbon dioxide concentrations are another important factor with a direct

physiological effect on fungal biomass. Carbon dioxide stimulates the growth of

mycelium because of its role in anaplerotic reactions in which carbon fixation occurs for

the formation of intermediate compounds in the TCA cycle, and in the formation of

purines, pyrimidines, and fatty acids (Maheshwari et al., 2000; Carlile et al., 2001). In A. bisporus vegetative growth, mushroom producers try to maintain carbon dioxide levels

between 5000-7000 ppm within mushroom production chambers. Once the mushroom

fungus is ready to fruit, carbon dioxide levels are reduced to 900-1200 ppm.

For mushroom production on composted substrates, the typical moisture content

of the substrate should be 63-68%. Moisture contents out of the optimal range hinders

the vegetative growth of the mushroom fungus. Consequently affecting the yield of

mushrooms. Flegg (1985) states that vegetative growth of the mushroom fungus is best at

moisture contents ranging from 55% to 70% in compost-based substrates. Wetter

substrates packed less densely were characterized with better growth and this was

assumed to be due to better aeration (Flegg, 1985). For non-composted substrates,

146 Mamiro (2007) observed that substrate moisture contents ranging between 55-60%

produced the highest yield of mushroom for a non-composted substrate composed

primarily of oak sawdust or of a combination of oak sawdust substrate with an additional

amendment of spent mushroom compost (SMC), the waste stream that is generated from

compost-based substrate. The highest mushroom yield of 14.5 kg/m2 with a

corresponding bioefficiency of 59.1% was obtained from a substrate mixture of oak

sawdust and spent mushroom compost with a moisture content of 55% (Mamiro 2007).

In mushroom science, one of the parameters for estimating the adequacy of

substrate moisture content or formulation is hyphal extension rate (Sanchez and Royse,

2001; Mamiro 2007). Higher hyphal extension rates are in most cases desired, but this

method is not suitable to directly quantify the amount of biomass being generated in the

substrate and only gives limited indication on whether or not the substrate is suitable for

mushroom production.

To estimate the amount of biomass and determine whether this substrate is

adequate for mushroom production, one must take into account the vegetative growth of

the mushroom fungus and ultimately the effect on mushroom yield. Direct estimation of

fungal biomass in solid-state fermentation (complete recovery of biomass) is not possible

(Raimbault, 1998). However, some studies have suggested the use of membrane culture

(Mitchell et al., 1991; Nagel et al., 2000) or the use of scanning electron microscopy

(Raimbault, 1998). In the case of membrane culture, total biomass is removed as the membrane is peeled off the underlying substrate. In reality, this cannot be applied to commercial SSF. Currently, the most widely used methods for fungal biomass estimation are indirect methods: respirometric measurement, quantification of extracellular enzymes,

147 biomass component estimation (glucoasmine, nucleic acids, protein content, and

ergosterol).

Aerobic microbial activity is associated with oxygen uptake and carbon dioxide

evolution. Hence, the oxygen uptake and carbon dioxide are growth associated (not in the

case of endogenous respiration) and can be used for the estimation of biomass

(Raimbault, 1998) and in this case for A. bisporus biomass estimation. Manometric

methods (pressure drop) is one way of measuring respiration rate, whereby a pressure

sensor records the pressure drop inside a sealed vessel containing a biomass sample. The biomass consumes the oxygen and releases carbon dioxide that is subsequently stripped

from the head-space by an absorbent leading to pressure drop inside the vessel. There are

few reports of poor or limited growth of fungi because of carbon dioxide deficiencies in

cultures in which carbon dioxide has been absorbed by alkaline propagules (Carlile et al.,

2001). Another method of estimating respiration rate is through the use of online or

continuous measuring systems in which inlet and outlet air are measured simultaneously

for oxygen, and carbon dioxide.

For this study, rye grain-based substrates with different moisture contents were

evaluated for OUR, linear extension rate, mushroom yield, and substrate bioefficiency.

This information will provide some insight upon the range of moisture contents for rye

grain-based substrates that can be used by mushroom producers.

9.3. Methods

9.3.1. Fungal Cultures and Substrate Materials

Rye grain spawn cultures of Agaricus bisporus (MC 459) were obtained from the

Mushroom Spawn lab (Penn State University), and millet commercial grain spawn was

148 obtained from Mushroom Test and Demonstration Facility (Amycel Maxx, Avondale,

PA). All fungal cultures of A. bisporus were stored at 5°C until use. The substrates used

in this study were primarily composed of rye grains that were supplemented with

Promycel Target® (SpawnMate, Watsonville, CA) when transferred to trays.

9.3.2. Substrate Preparation for OUR Measurements and Mushroom Fruiting

Rye grains, water, CaCO3 and CaSO4 (Fisher Scientific, Hampton, NH) were

added to respirometer bottles (Oxitop®-WTW, Giessen, Germany) for the OUR

measurements or to 1000 ml Erlenmeyer flasks for the mushroom fruiting experiments to

achieve the following moisture contents 45, 50, 55, 60, 65, and 70%. The ratio of calcium

carbonate and calcium sulfate to grain was maintained at 1:1:50, respectively. The total

mass of substrate in each bottle/flask was set at 450 g not including the CaCO3 and

CaSO4 amendment. Table 9.1 provides a summary of the mixtures for each moisture

treatments.

Table 9.1. Substrate formulations for the different moisture treatments Rye Grain Water CaCO CaSO Treatment 3 4 (g) (g) (g) (g) 45% 275 175 5.5 5.5 50% 250 200 5 5 55% 225 225 4.5 4.5 60% 200 250 4 4 65% 175 275 3.5 3.5 70% 150 300 3 3

The bottle and flasks were sealed with cotton wool plugs and sterilized in an

autoclave for 45 min (121oC- 103.4 kPa). The bottles and flasks were transferred to a laminar flow hood and kept there overnight. The following day, each bottle or flask was aseptically inoculated with 8 g of A. bisporus rye grain spawn under the laminar flow

hood and transferred to an incubator in which temperature was maintained at 22oC.

149 9.3.3. Procedure for OUR Measurement

Oxygen uptake rate of the different treatments was measured using the Oxitop® system (WTW, Giessen, Germany). The system is composed of a 1.209 L glass bottle in which a biomass sample and a bottle sealing cap composed of a receptacle for a carbon dioxide absorbent (typically NaOH), rubber gaskets, and a pressure sensor head that transmits the enclosed bottle pressure reading to a controller. The Oxitop® system has been successfully used in several studies for the measurment of compost and compost bulking material OURs (Sadaka et al., 2004; Ahn et al., 2005).

When the OUR of millet grain spawn was measured, the grain spawn was added to pre-sterilized Oxitop® (WTW, Giessen, Germany) bottles and the measurement was taken for a 24 hr duration. A sample of the grain spawn was taken for dry matter and ash content determination.

OUR measurements of culture growing on substrates with different moisture contents were done over a period of 32 days with each individual measurement taking 24 hrs as stated earlier. Ash and dry matter content were measured by calculating the individual contributions of each of the components of the substrate, at the beginning of the test (before inoculation) and at the end of the experiment (after the 32 day period) for ash and dry matter analysis. This minimized the risk of contamination and reduced the amount of sheer stress on the actively growing cultures. The change in dry matter and moisture was assumed to be linear and the results were integrated into the calculations for the OUR.

150 9.3.4. Oxygen Uptake Rate (OUR) Calculation

The oxygen uptake rate was calculated based on the pressure drop in the apparatus. The pressure drop is induced by the oxygen consumption by the living biomass. The pressure drop data was converted into OUR by using equation 9.1.

⎛ hPa⎞ ⎛ Pa ⎞ 3 slope⎜ ⎟ ×100⎜ ⎟ ×Vgas()m × 32()g/moleO2 ×1000()mg/g ×1440() min day ⎛ mgO ⎞ ⎝ min⎠ ⎝ hPa⎠ OUR⎜ 2 ⎟ = Eq. 9.1 ⎝ gVSday⎠ ⎛ J ⎞ 8.314⎜ ⎟ × TK()×Wg()× (1 − MC) ×VS ⎝ moleK ⎠ where

slope = slope of the pressure drop curve

Vgas = volume of gas occupying the Oxitop vessel

T = temperature

W = weight of sample

MC = moisture content wet basis

VS = volatile solids content

the remaining numbers are conversion factors.

To calculate the volume of gas inside each bottle the following equation was used as shown in equation 9.2.

3 Vgas(m ) = Vbottle −Vsubstrate + Vsubstrate ×εa Eq. 9.2

where

3 Vsubstrate = volume occupied by substrate (m )

εa = air filled porosity (%)

Air filled porosity was calculated using equation 9.3 as described by Richard et al.

(2002).

151 ⎛ MC DM ×VS DM × (1−VS)⎞ εa =1− ρwb ×⎜ + + ⎟ Eq. 9.3 ⎝ ρw ρOM ρash ⎠

where

3 ρwb= bulk density of material (kg/m )

3 3 ρOM= 1.6 x 10 (kg/m )

3 3 ρash= 2.5 x 10 (kg/m )

MC= moisture content

DM= dry matter content

9.3.5. Dry Matter and Ash Measurements

For all materials (grain, substrate, casing, and perlite), 10 g samples were placed

in ceramic crucibles and then placed in an oven at 105 oC for 24 hrs. The weight was

measured and the dry matter content was calculated. The crucibles were then placed in a

muffle furnace at 550 oC for 9 hrs. The residual weight was measured with ash and

volatile solids content were calculated.

9.3.6. Linear Extension Rate Measurements

Test tubes and caps were sterilized for 20 min in an autoclave and allowed to cool

under a laminar flow hood. Once cooled, two particles of rye grain spawn were added to

each individual test tube. Next, pre-sterilized and cooled rye grain substrate prepared in

1000 ml Erlenmeyer flasks at the different moisture levels (45, 50, 55, 60, 65, 70%) were

filled to the top of the test tubes. The test tubes were capped and placed in an incubator at

24 oC. Hyphal extensions were measured using a digital caliper (Digimatic caliper,

Mitutoyo, Kanagawa, Japan). Hyphal extension rates were calculated using the slope of the curves generated using Microsoft Excel (Microsoft Corporation, Redmond, WA).

152 9.3.7. Experimental Set-up

The first experiment tested whether substrate mass (millet grain spawn) within the

Oxitop® and the NaOH mass as CO2 absorbent had a direct effect on OUR. Three different levels of millet grain spawn (50, 150, 250 g) and NaOH (1, 1.5, 2.5 g) masses were undertaken and replicated thrice.

The second set of experiments measured the OUR for different rye-grain substrate moisture contents (45, 50, 55, 60, 65, and 70%) and tested whether measurement frequency (every 2 days or every 8 days) had an effect on OUR. The amount of NaOH added to each bottle cap receptacal was 1.5 g. Each measurement lasted for 32 days and this experiment was replicated twice. Mushroom yield from the substrate in the Oxitop® bottles was determined by adding 280 g of rye grain substrate with 14 g of Promycel

Target® to sterilized Ziploc containers with 200 ml of wetted perlite and cased with a mixture of mushroom casing containing activated carbon (10% by volume). Furthermore, rye-grain substrate at the different moisture levels were measured for hyphal extension rate.

The last set of experiments increased the substrate mass to 800 g to observe if any changes in mushroom yield and substrate bioefficiency for a rye grain substrate with the different moisture contents occur with scale-up. These substrates were prepared in

Erlenmeyer flaks and the procedure for transfer to trays was comparable to the methodology presented in Bechara et al. (2006b). The substrate was supplemented with

5% Promycel Target® at casing, and treatments were replicated 4 times.

153 9.4. Results

9.4.1. OUR Measurements as affected by the Mass of Substrate and NaOH Pellets

These treatments tested whether OUR was dependent on the interaction effect of

substrate mass x NaOH mass. The results indicated that the only significant effect on

OUR was the substrate mass which means that any of three NaOH masses were sufficient in absorbing all of the carbon dioxide generated by the biomasss growing in the substrate.

The main effect NaOH mass and the interaction effect, as stated earlier, were not

significant (p>0.05). Table 9.2 summarizes the observed OURs for the different

treatments.

Table 9.2. Observed OURs for treatments in which substrate mass and NaOH mass were varied.

NaOH OUR1,2 Millet Grain Spawn (g) (mg O /g VS day) (g) 2 50 1 5.57 ± 0.21a 50 2.5 4.76 ± 0.50a,b 50 4.5 4.89 ± 0.61a,b 150 1 4.36 ± 0.28a,b 150 2.5 4.55 ± 0.30a,b 150 4.5 4.21 ± 0.78a,b 250 1 3.97 ± 0.22b 250 2.5 4.41 ± 0.63a,b 250 4.5 4.44 ± 0.40a,b 1 Data presented with standard deviations 2 Treatments followed by similar letters are not significantly different (p<0.05, is significant)

The highest OUR was generally observed for the 50 g substrate treatments,

whereas the lowest were observed for the 250 g treatments as shown in Figure 9.1.

154

5.1

5.0

4.9

4.8

4.7 / gVS day) / 2

4.6

4.5

4.4 OUR (mg O

4.3

50 150 250

Figure 9.1. OUR observations as affected by the mass of the substrate with the red-line indicating OUR when all the observations are average.

9.4.2. OUR Readings as Affected by Substrate Moisture Content

The effect of moisture content of the rye grain substrate on the OUR of A. bisporus was statistically significant (p<0.05), and the highest OUR was observed for the

65% treatment (4.88 mg O2/g VS day). The highest OUR observed for the different

moisture treatments is shown in Table 9.3.

155 Table 9.3. Peak OUR for A. bisporus growing on a rye grain substrate with different moisture contents.

Peak OUR Moisture Treatment Day Observed (mg O /g VS day) (%) 2 70 4.34 ± 2.16 12 65 4.88 ± 2.03 24 60 3.85 ± 0.46 28 55 3.17 ± 0.34 28 50 2.50 ± 0.06 28 45 1.96 ± 0.73 28

After day 18, the OUR of the 70% treatment, although the highest until then, decreased as compared to all other treatments. In addition, an uncolonized zone (no fungal mycelium) was observed at the bottom of the bottles for the 70% treatments, whereas the top part of the substrate was colonized with the mycelium. This effect is thought to be due to oxygen limitations in the bottom part of the flasks. Furthermore, this decrease in OUR is also observed for the 65% treatment beyond day 24, and the 50% treatment beyond 30th day. However, none of the other treatments developed an uncolonized zone as did the 70% treatment. For all other treatments, the OUR remained stable. Figure 9.2 depicts the change in OUR for the different moisture levels over a period of 32 days.

156 6

5 y 4 70% 65% 60% 3 55% 50% 2 45% OUR (mg O2/g VS da

1

0 2 4 6 8 10 12 14 16 18 20 22 24 26 28 30 32 Day

Figure 9.2. OUR of A. bisporus growing on a rye grain substrate with different substrate moisture contents treatments with rye grain spawn.

Figure 9.3 depicts the different treatments growing in the Oxitop® bottles. For, the 70% moisture level, there is a clear zone of uncolonized grains below a layer of colonized grains.

157

Figure 9.3. Different moisture level treatments for a rye grain-based substrate inside the Oxtop® bottles.

The calculated rye grain substrate porosity was lowest for the 70% treatment at the beginning and at the end of the OUR measurements and was highest for the 45% treatment. The reason the porosity was so low for the 70% treatment was that the grains lost their structural integrity. Furthermore, whenever the flasks were shaken to fragment the mycelium and disperse the biomass to other portions of the substrate, grain structural integrity worsened. The increase in porosity at the end of the OUR measurements was due to the growth of the fungal biomass. Whenever the substrate was shaken the fungal matt that developed over the substrate and between the rye grains was sheered and

158 different sized particles formed. Figure 9.4 depicts the change in porosity for a rye grain

substrate at different moisture levels.

0.6

0.5

0.4 ) % (

y final 0.3 initial

Porosit 0.2

0.1

0 70 65 60 55 50 45

Moisture Levels (%)

Figure 9.4. Change in porosity for the different rye grain substrate at different moisture levels at the beginning and at the end of the OUR measurement that spanned 32 days.

Mushroom yield and substrate bioefficiency from the Oxitop® bottles varied significantly (p<0.05). The peak mushroom yield and substrate bioefficiency of 232 g and

174% were observed for the 55% treatment. The 70% moisture treatment did not yield mushrooms. Figure 9.5 depicts observed yields and substrate bioefficiencies for the different treatments.

159 300

250

200

150

and BE (%) 100 Mushroom Yield (g)

50

0 45 50 55 60 65 70

Moisture Levels (%) Figure 9.5. Mushroom yield and substrate bioefficiency for rye grain substrates from the Oxitop® bottles with different moisture levels supplemented with 14 g of Promycel Target®.

9.4.3. OUR Readings as Affected by Measuring Frequency

The OUR of A. bisporus was influenced by the measuring frequency for most

moisture levels except for the 70% and 45% treatment. For the 70% treatment, the day 32

measurement of the 2 day measuring frequency was 2.91 mg O2/g VS day, which was far

greater than the OUR of 8 day measuring frequency (0.88 mg O2/g VS day). The reason

why this occurred may have been due to contamination with anaerobic organisms. In

cases where the measuring frequency was significant, the 8 day frequency exhibited

higher OUR values than the 2 day frequency measurement treatment. An additional

observation was made that pressure drop leveling off (i.e. total anaerobic conditions) was

attained half-way (around 10-15 days) through the duration of the experiment for the

70% and 65% moisture levels, and gradually extended to the lower moisture levels as the

160 experiment proceeded with the exception of the 45% moisture level treatment. This is

why the 45% treatment did not exhibit any differences in OUR with measuring frequency. Table 9.4 provides a summary of the data for the different treatments along with the p-values associated with the statistical analysis.

Table 9.4. OUR of A. bisporus influenced by frequency of OUR measurements growing on a rye grain substrate with different moisture contents. OUR for 2 day Frequency OUR for 8 day Frequency (mg O2/g VS day) (mg O2/g VS day) Moisture Day Day Day Day Day Day Day Day p-value1 (%) 8 16 24 32 8 16 24 32 70 2.82 4.75 3.58 0.88 3.89 4.66 2.89 2.91 0.675 65 1.86 3.52 4.09 3.47 3.61 6.39 5.74 5.28 0.002 60 0.75 2.68 3.33 3.51 1.65 3.87 4.71 5.22 0.007 55 0.90 2.28 2.79 2.84 1.08 3.11 3.61 3.79 0.010 50 0.56 1.56 2.07 1.43 0.66 2.16 2.51 2.72 0.041 45 0.32 0.92 1.37 1.52 0.27 1.09 1.51 1.81 0.366 1values within a row were statistically analyzed using GLM and p<0.05 indicates there is a significant difference between the two measurement frequencies.

9.4.4. Mycelium Extension Rate

The mycelium extension rate varied among the different moisture treatments. The

highest extension rate was observed for the 70% rye grain treatment, whereas the lowest

was observed for the 45% treatments. In addition, it took treatments with lower moisture

contents a greater time before growth started occurring. For the 45% moisture treatment,

hyphal extension started occurring 14 days after inoculation, whereas it occurred 5 days

after inoculation for the 70% and 65% treatments. Furthermore, the lower moisture

contents (45-55%), especially the 45% treatment were more prone to exhibit no growth,

as not all of the replications for those three moisture levels grew. Figure 9.6 shows the

difference in extension rate and lag phase duration for the different treatments.

161 10

9

8

7

6 Hypal Ext 5 Lag Phase 4

3 Lag Phase Duration (days) (days) Duration Lag Phase 2 Hyphal Extension Rate (mm/day) and Rate (mm/day) Hyphal Extension 1

0 70 65 60 55 50 45

Moisture Levels (%)

Figure 9.6. Hyphal extension rate and lag phase duration for the different rye grain substrate with different moisture contents.

Furthermore, a visual observation of the treatments showed that the density of the

mycelium was strikingly lower for treatments with lower moisture contents than for

treatments with the higher moisture content. Figure 9.7 shows the different treatments

depicting the differences in mycelium density.

162

Figure 9.7. Hyphal growth of A. bisporus on the rye grain substrate with different moisture contents.

9.4.5. Yield and Substrate Bioefficiency as Affected by Substrate Moisture Content

Mushroom yield and bioefficiency varied over the range of moisture treatments.

The statistical analysis indicated that the effect of moisture content on bioefficiency was significant (p<0.05) but was not significant for yield (p>0.05). Large variation in yield

for the 45% and 70% coupled with the fact that mushroom yield for the moisture ranges

between 50-65 % were comparable on average had rendered the effect of moisture level

on yield not significant. Furthermore, two out of the four replications for both the 45%

and 70% treatment failed to produce mushrooms, which was not observed for the other

four moisture treatment levels. The lowest yield was observed for the 70% treatment

followed by the 45% treatment, yielding 163 g and 223 g, respectively. It is important to

mention, however, that two replicates for both treatments failed to produce mushrooms.

Although not statistically significant, the highest mushroom yield was observed for the

60% treatment.

163 The highest bioefficiency was observed for the 65% moisture treatment (162%)

and the lowest was observed for the 45% treatment. Figure 9.8 depicts the yield and

bioefficiency of the different moisture treatments.

600

500

400

300 Yield and BE (%) BE Mushroom Yield (g) 200

100

0 45 50 55 60 65 70

Moisture Levels (%)

Figure 9.8. Mushroom yield and substrate bioefficiency for different grain-based substrates with the grain portion varying in moisture content.

9.5. Discussion

Moisture content of the substrate, as in any solid state fermentation process, is a critical parameter that mitigates the growth of the mushroom fungus. In this study, the moisture content of a rye grain substrate ranging from 45-70% was shown to have a direct influence on OUR, mycelium extension rate, mushroom yield, and substrate bioefficiency. However, a distinction must be drawn between the vegetative stage of A.

bisporus and the mushroom production stage.

164 First and foremost, it is important to note that OUR of A. bisporus increased with

increasing moisture contents. However, it was observed that the measuring process

impacted the growth of the fungal biomass. This observation is based on differences

between the OUR measurement frequency experiment, whereby the OURs were lower

for the set of treatments that were measured more frequently. A more suitable method of measuring OUR would be a continuous system in which inflow and outflow gas concentrations are measured continuously.

The greater the hyphal extension rate, the shorter is the spawn-run duration. This is desirable because the mushroom production process time is reduced. Furthermore, the density of the mycelium was far greater for the higher substrate moisture contents

compared to the lower moisture contents, similar to what Shroeder and Schisler (1981)

observed for compost-based substrates. However, Gerrits (1971) observed that moisture

contents for compost-based substrates greater than 72% reduced mycelium growth.

Mamiro (2007) observed that hyphal extension rates decreased for non-composted

substrate moisture contents ranging between 60-65%, but this was not observed for rye-

grain substrate tested herein. This is certainly a manifestation of dependency of substrate

moisture contents on substrate formulations and compositions, and that optimal moisture

contents for different mushroom substrate types is variable.

However, for rye-grain substrates, the OUR data and the hyphal extension rate

appear to go hand in hand; the higher moisture contents have a higher OUR and a larger

hyphal extension rate compared to the drier substrate. However, higher OUR translates to

a greater consumption of the carbon source at a stage in which the A. bisporus is not

producing any mushrooms. Hence, it would be important to try to find a suitable moisture

165 content at which the vegetative growth of the mushroom fungus is rapid and mushroom

production is high. In small scale experiments, the 55% moisture level produced the

highest yield, whereas for the scale-up treatments yield for moisture levels ranging from

50-65% were comparable. The reason for the difference is due to the duration of the OUR

measurements which spanned 32 days. In fact, by day 18 the treatments with 55-65%

moisture levels were fully colonized. In the scale-up experiment, as soon as the substrates

were fully colonized by the mushroom fungus they were immediately transferred,

minimizing the loss of dry matter to vegetative growth, whereas the smaller OUR experiments consumed most of the energy reserves within the grains. Although mushroom yield is important, the comparison of yield from substrates having different moisture and dry matter contents may not be an adequate interpretation of the data.

Normalizing the data based on dry matter content i.e. substrate bioefficiency is more accurate. In the small-scale experiments, the highest bioefficiency was observed for 55% moisture level, where the highest bioefficiency in scale-up experiments was observed for the 65% moisture level. Hence, 60-65% moisture seems to be the best moisture level treatment.

One factor of primary concern is the handling of the grain substrate during processing. As the moisture content increases, an observed increase in substrate stickiness (decrease in flowability) is observed. This is in part due to the leaching and solubilization of starch granules (gelatinization). Furthermore, for the higher moisture contents, the grain integrity becomes compromised and shaking the flasks induces a total structural collapse of individual grains rendering the substrate “doughy”. All of these observations have lead mushroom spawn producers to operate at low spawn substrate

166 moisture contents i.e. 48-55%. These data are based on grain moisture content of the final product, as the initial moisture content is proprietary. It appears, as although substrate moisture contents ranging between 55-65% are adequate for rye grain substrate, moisture contents of 45% and 70% are not because of increased failure of growth at the lower and higher end moisture levels. More work is needed to determine whether the same moisture content ranges hold for different types of grain-based substrates.

9.6. Conclusions

The moisture content of rye-grain based mushroom substrates had a direct effect on OUR, hyphal extension rate, mushroom yield, and substrate bioefficiency. OURs and hyphal extension rates increased with increasing moisture contents with highest OUR of

4.88 mg O2/g VS day observed for the 65% treatment. OUR measurements using

pressure sensor methods appears to negatively impact the growth of the mushroom

fungus and it is recommended that continuous respirometric measurement methods are

used to ensure that aerobic conditions are maintained throughout the test duration, and

that the stimulating effect of carbon dioxide on fungal biomass growth is maintained at optimal levels. Mushroom yield and substrate bioefficiency were highest for substrate moisture contents ranging from 50-65%, whereas 45% and 70% moisture contents were

not suitable.

167 Chapter 10

Two Alternative Agaricus bisporus Mushroom Production Systems Using Grain- based Substrates

10.1 Abstract

Two systems for Agaricus bisporus (button mushroom) production are proposed as alternatives to the traditional environmentally problematic mushroom production

system that relies on composting of plant and animal organic matter. Each system

involves the use of cereal grains and lesser ingredients as a mushroom substrate. The first

system, called the “Satellite Mushroom Production System” (SMPS), proposes the use of

commercial grain spawn, the vehicle typically used to inoculate composted substrates,

supplemented with high protein delayed-release supplements as a non-composted

mushroom substrate. The second system called “ Complete On-site Mushroom

Production System” (COMPS) consists of producing mushrooms on sterilized grains

supplemented with oilseeds and lesser ingredients. In this system, an aseptic processing

system would be located on-site at the mushroom production facility to convert the raw

materials into a grain-based substrate. In this research, the highest yield of mushrooms

for the commercial grain spawn substrate supplemented with delayed-release

supplements was 14.28 kg/m2, whereas yield from substrates composed of cereal grains

and oilseeds was 21.3 kg/m2. Finally, a cost model was developed for the proposed grain-

based mushroom production systems. Based on this model, the direct costs (ingredients,

labor, operating costs) to produce mushrooms using the SMPS was $4.00 /kg mushroom,

whereas that of the COMPS was $1.40 /kg mushrooms. Mushrooms produced on

168 compost-based substrates cost around $0.70-0.80/ kg mushrooms which is lower than for

the grain-based system.

10.2. Introduction

Agaricus bisporus (J.E. Lange) Imbach, the button mushroom, is produced worldwide and available strain types are: white, off-white, hybrid, and brown strains

(Fritsche and Sonnenberg, 1988). In 2005-2006 total Agaricus mushroom sales in the

USA were valued at $841 million with a corresponding crop size of 376,482 metric tons

(USDA, 2005-2006). The brown mushrooms (Portabello and Crimini) varieties sales

were $152 million with a total crop size of 53,070 metric tons (USDA, 2005-2006).

Mushroom producers received $2.13/kg for white varieties and $2.86/kg for brown

varieties. The problems facing Agaricus bisporus mushroom producers are related to substrate preparation and spent mushroom compost disposal.

The most widely used substrate for A. bisporus mushroom production is

composed of plant and animal organic matter that has been composted. Sinden and

Hauser (1950) developed the two-phase composting process that begins with an outdoor

composting phase followed by an indoor phase, during which temperature is closely controlled. Composting within the mushroom production process is a lengthy (may take

up to three weeks), malodorous, and labor-intensive step (Derikx et al., 1990; Bechara et

al., 2006a). Furthermore, after the substrate no longer supports economically viable crop

yields (spent mushroom compost – SMC), the substrate is pasteurized, removed from the

production chambers, and usually applied to fields to weather for 2-3 years (Heinemann

et al., 2003). The resumption of odor production is observed because SMC has not been

biologically stabilized. Hence, the degradation of organic matter (composting) continues

169 and odor production resumes. Disposal by land application is limited by the fact that

SMC is rich in salts and nutrients that are capable of impairing the quality of surface water and groundwater (Williams et al., 2001; Guo et al., 2001). In Pennsylvania, permits are required to land-apply SMC and this has become a problem because of the large amount of SMC estimated at 580,000 m3/ year (Heinemann et al., 2003; Beyer 2006).

Producing A. bisporus mushrooms on non-composted substrates is possible, and interest in non-composted substrates has re-emerged because of the negative impact mushroom operations are having on neighboring residents. Till (1962) refined a substrate composed of chopped and ground straw, white peat, calcium carbonate, cottonseed meal and soybean meal, and this was commonly known as Till’s substrate. After mixing, moistening, and sterilizing the substrate, it was inoculated with A. bisporus. Till (1962) observed that the colonization phase of non-composted substrate was longer compared to that of composted substrate, but yield in kg/ton of substrate was high. This process was never adopted because of high operating costs. Huhnke and Von Sengbush (1968) simplified the Till process by replacing sterilization with pasteurization. The substrate was steamed and then inoculated with a suspension of microorganisms from compost.

However, the results were inconsistent.

Smith and Hayes (1972) grew A. bisporus on inert sphagnum peat moss with nutrients supplied in a solid form. The maximum yield obtained was 16.7 g, which was higher than the liquid nutrient solution treatment (7.1 g).

A method of producing A. bisporus fruiting bodies from cased grain spawn was developed by San Antonio (1971). The grain spawn formulation was composed of rye grains mixed with calcium carbonate and de-ionized water (20 g rye grain/0.4 g

170 CaCO3/20 ml di-water). The mixture was sterilized (121°C for 60 min), cooled and

inoculated with 10-20 grains of spawn. Once A. bisporus colonized the entire substrate, a

layer of pasteurized casing was spread over the colonized grains. As mycelium reached

the surface, aeration was increased and mushrooms were produced. Although yield of

mushrooms was not presented, San Antonio (1971) concluded that the quantity of

mushrooms produced was comparable to that obtained from conventional compost. Mee

(1978) was able to obtain “good quality” mushrooms on a non- composted substrate

formulation composed of cold manure, sphagnum peat moss, and gypsum.

Sanchez and Royse (2001) refined a substrate composed of pasteurized mixture of

oak sawdust, millet, rye, peat, alfalfa meal, soybean flower, wheat bran, and calcium

carbonate and found that it was suitable for the production of A. bisporus (brown

Portobello). The maximum biological efficiency (fresh weight of mushrooms divided by the dry weight of the substrate) achieved was 77.1% and mushroom yield was 31.4 kg/m2. Reported mushroom yield was higher than in traditional composted substrates but

bioefficiency values were low. Schisler (1982) states that observed bioefficiencies of

80% or higher for composted substrates are considered good. Mamiro et al., (2007)

showed that a 1/1 ratio of non-composted substrate (similar to Sanchez and Royse) and

SMC produced a mushroom yield of 27.2 kg/m2 with a corresponding bioefficiency of

144%.

Bechara et al., (2006a) showed that a non-composted grain substrate composed of

millet grains, perlite, and calcium carbonate produced mushroom yields comparable to

composted substrates (8.7 kg/m2 and 7.7 kg/m2, respectively). Replacing the millet grains

with commercial rye grain spawn yielded less mushrooms (5.3 kg/m2). However, adding

171 delayed-release supplements to the commercial grain spawn increased yield from 1.4

kg/m2 to 7.6 kg/m2, and the addition of an underlying layer of perlite below the substrate

further increased mushroom yield to 13 kg/m2 (Bechara et al., 2005b). The highest yield

obtained from a commercial grain spawn substrate was 14.28 kg/m2, and the

corresponding substrate bioefficiency was 177%. A sterilized mixture of cereal grains

and cracked roasted soybean colonized with A. bisporus and then supplemented with

delayed-release supplements (Full House –S41) produced a mushroom yield of 16.9

kg/m2 compared to 8.7 kg/m2 for a substrate composed of commercial millet grain spawn

supplemented with a the same delayed-release supplement (Bechara et al., 2006b).

Further refinement of the cereal grain and oilseed substrate increased mushroom yield to

21.7 kg/m2 with a corresponding bioefficiency of 240% (unpublished data).

The objective of this study is to present two theoretical mushroom production

systems that use non-composted grain-based substrates as alternatives to the commercial

compost-based system. Furthermore, a theoretical design for a continuous aseptic

processing system for grain substrate processing to be used in two of these systems is

presented in detail. In addition, a brief discussion of the steps in mushroom production

beyond the substrate preparation phase and a preliminary cost analysis for each process is

presented.

10.2.1. Steps in Agaricus bisporus Mushroom Production

The first step in a mushroom production process is substrate preparation. Steps for

mushroom production beyond the substrate preparation phase are adopted universally by

A. bisporus mushroom producers. In all cases, the substrate is prepared, then inoculated with grain spawn and placed in tray reactors. Temperature (22-24 ◦C), humidity (85-

172 90%), and carbon dioxide levels (5000 ppm) are monitored and controlled within acceptable ranges conducive to the vegetative growth of the mushroom fungus.

After the substrate is entirely colonized, a top-dressing, termed “mushroom casing”, is spread over the substrate surface. Mushroom casing can be added immediately after inoculation, resulting in earlier mushroom production. The casing layer is necessary for A. bisporus fruiting. Environment temperature (16 ◦C), relative humidity (75-80%) and carbon dioxide concentration (800-1200 ppm) must be controlled within specific ranges that promote A. bisporus fruiting. Mushrooms are then manually harvested until additional yield is no longer economically viable (typically 2-3 weeks). Finally, mushroom tray reactors are steamed to inactivate pathogens that may be present, and the residual material is called “spent mushroom compost” (SMC). The SMC is removed from the mushroom production facility and stacked outdoors. Figure 10.1 summarizes the steps in Agaricus sp. mushroom production beyond the substrate preparation phase.

173

Figure 10.1. Overview of steps in Agaricus sp. mushroom production beyond the substrate preparation phase.

174 10.2.2. Factors Influencing Mushroom Yield in Grain-based Substrates

Previous work in grain-based substrates has identified key factors that have a significant effect on mushroom yield. Mushroom yield for commercial grain spawn substrates is influenced by:

• amount and type of delayed-release supplement (Bechara et al., 2005a)

• adding activated carbon to sterilized casing and non-sterilized casing

(unpublished data).

• using an underlying layer of water-holding materials (Bechara et al., 2005b)

• commercial grain spawn age (unpublished data)

Additionally, for treatments using a grain/oilseed substrate, mushroom yield was influenced by

• type and rate of oilseed added to the substrate (Bechara et al., 2006b)

• addition of delayed-release supplements at casing (Bechara et al., 2006b).

• type and rate of oilseed added to the substrate (Bechara et al., 2006b)

• supplementation with delayed-release supplements at casing (Bechara et al.,

2006b).

• type of cereal grain used and pre-incubating the grain substrate with Scytalidium

thermophilum- a mushroom compost thermophile.

Figure 10.2 depicts a tray with mushroom fruiting on a substrate composed of

grain/oilseeds supplemented with a commercial delayed-release supplement added over a layer of perlite.

175

Figure 10.2. A tray with mushroom fruiting from a substrate composed of millet grains and cracked soybean supplemented with Promycel Target® (delayed-release supplement) with an underlying layer of perlite.

10.3. Description of Alternative Mushroom Production System In this section, two theoretical mushroom production systems are described in

detail. The first process is called SMPS, and the second is called COMPS.

10.3.1 Satellite Mushroom Production System

The SMPS is a system in which commercial mushroom grain spawn and delayed-

release nutrient supplements are used as the basal substrate for mushroom production.

Mushroom producers would purchase fully colonized grain spawn, delayed-release

supplements, peat moss, and lime from appropriate suppliers. The substrate components

(commercial grain spawn + delayed-release supplements) are mixed on-site and added to

mushroom trays containing an underlying layer of perlite or other water-holding

materials, and then cased with peat-based mushroom casing. Equipment necessary to

perform all of these tasks is basically the same as that used for compost-based substrates,

176 is found on traditional commercial mushroom production facilities, and can be purchased

from equipment suppliers. The mushroom trays are then transported to environmentally

controlled chambers and cropping proceeds as in the traditional system. Figure 10.3

summarizes the different steps involved in the SMPS.

Figure 10.3. Overview of steps in the SMPS for Agaricus sp. mushroom production using a substrate of commercial grain spawn and delayed-release supplements.

10.3.2. Complete On-site Mushroom Production System

The COMPS differs from the SMPS in that the grain-based substrate is processed using a continuous aseptic processing system and colonized with the mushroom fungus

on-site. The aseptic processing system would be formed of four sections that are

combined to provide continuous production of grain-based mushroom. The aseptic

processing system described is based on segmented-flow technology, a Penn State

patented technology (Walker, 2002). The four sections are described both in Table 10.1

and Figure 10.4.

177 Table 10.1. Description of individual processes occurring in the continuous aseptic processing unit for A. bisporus production on non-composted grain-based substrates in the COMPS. Process Occurring in Section Grain moisture content control. Cereal Section I grains are heated in the presence of excess water to achieve targeted moisture content Cereal grains and other additives (oilseeds Section II etc) are sterilized using pressurized steam Grain substrate components are cooled and Section III mixed aseptically in an air pressurized paddle mixer Cooled grain substrate is inoculated with Section IV the mushroom fungus and then aseptically added to sterile filter-patch bags

Section I is composed of an “undershot” conveyor system with a belt that has

dividers or barriers that segment the flow of grains into specific control volumes to

approximate a plug-flow residence time distribution. The conveyor system is driven by a

variable speed drive motor so the operator can adjust residence time to achieve the

desired grain moisture content.

Section II is composed of a flight conveyor system constructed within a steam

pressurized sanitary tube and three rotary valves. The role of the rotary valves is to

transport the substrate materials while preventing steam loss and air entry into Section II.

Two of the rotary air-lock valves are located at the two ends of Section II. The third

rotary valve is positioned over the conveyor belt, upstream to the water/steam interface, with the purpose of adding “other” ingredients such as calcium carbonate, calcium sulfate, minerals, and oilseeds. The steam inlet is placed at the top end of Section II and the nozzle is directed towards the returning belt system to dislodge any residual substrate

material.

178 Section III is composed of a jacketed, air-pressurized paddle mixer with a cooling fluid filling the jacket. The need for a pressurized cooling section is to minimize water phase change within the grains that may destroy grain structure and endosperm integrity.

Finally, Section IV is a clean room in which the substrate enters from Section III via an airlock valve into a Y shaped junction. The one end is used to add the inoculum to the sterilized and cooled substrate. Figure 10.4 depicts a theoretical schematic of the continuous aseptic processing unit that could be used in the COMPS.

179

Figure 10.4. Four stages of the aseptic processing unit for non-composted grain-based substrate in the COMPS.

180 In summary, cereal grains and makeup water are added to the upper end of

Section I (as shown in Figure 10.4) where the grains are directly submerged in heated

water and conveyed through Section I. The moistened grains exit into Section II through

rotary airlock valve where the grains are conveyed belt out of the water and into a steam

environment. Once out of the water, oilseeds and other lesser ingredients are added onto

the conveyor belt over the wetted grains and all the materials are sterilized. From Section

II, the material is dropped onto another rotary air-lock valve and into Section III where the ingredients are mixed by the paddle mixer and cooled by conductive cooling. The cooled grain substrate exits through the opposite end of the paddle mixer and enters

Section IV via a rotary air-lock valve. In Section IV, the grain substrate is inoculated and

added to bags manually. After this Stage IV the bags are transported to environmentally

controlled chamber where A. bisporus proceeds to colonize the substrate. Once, colonized

the substrate is then transferred to trays and cropping occurs similarly to the “Satellite

Mushroom Production System”. Figure 10.5 provides a diagram of the different steps

involved in the COMPS.

181

Figure 10.5. Overview of steps COMPS for Agaricus sp. mushroom production using a substrate of cereal grains/oilseed substrate and delayed-release supplements.

10.3.3. Duration of Mushroom Production Process

The mushroom production systems are characterized by having different process times, whereby the longest process duration is found for the compost-based system and shortest for the SMPS. Most of the process time for the compost-based system is spent on preparing the substrate. The first stage duration (8 days) includes a pre-wetting the material followed by Phase I (7 days) and Phase II (6 days) composting.

For the COMPS, most of the process time is spent on spawn run because the mushroom fungus growth on the non-composted grain-based substrates is much slower.

The 28 day value for spawn-run duration is for grain-based substrates that have been pre- incubated with S. thermophilum. It is anticipated that increasing the inoculation

182 (spawning) rate and decreasing the inoculum size (increase inoculation sites) will eventually decrease the spawn run duration. Furthermore, adding casing inoculum to the casing material could further reduce the process by 2-3 days.

The SMPS has the shortest turn-around because most of the grain-based substrate

preparation and spawn-run occurs upstream from the mushroom farm. Therefore, what is

left is strictly the time it takes for A. bisporus to grow and fruit. Table 10.2 provides a

summary of the different process duration for the various mushroom production systems.

Table 10.2. Process durations for traditional compost-based system compared to the SMPS and COMPS. Mushroom Compost-based Satellite Production Complete On-site Production Phase System System Production System (days) (days) (days) Pre-wet 8 - - Phase I 7 - - Phase II 6 - - Spawn-run-Casing 14 - 28 Casing-Pinning 7 10 10 Pinning-Harvest 1 14 11 11 Harvest 1- Harvest 3 14 26 26 Total 70 47 75

10.4. Cost Model for Grain-based Mushroom Production System

Two cost models were developed to compare the production cost of mushrooms

($/kg) for the two grain-based systems, and to perform a sensitivity analysis. The models

only include direct costs such as substrate formulation, energy, water, electricity, and

labor costs. The production capacity for the mushroom farm is 20 000 kg of

mushrooms/30 days. Using this input, all the remaining inputs were calculated. The

amount of substrate, perlite, and casing were comparable to the experimental rates in the

studies. For the COMPS, the costs and yield were based on a millet-based substrate

183 supplemented with 5% Promycel Target® and yield was 21.7 kg/m2. For the SMPS, costs

and yield were made for a commercial grain spawn substrate supplemented with 5% S41

producing a yield of 14.2 kg/m2. For more details, Appendix C contains the cost model

spreasheets. Table 10.3 provides some of the inputs used in the both the SMPS and

COMPS cost model.

Table 10.3. Common parameters for the SMPS and COMPS cost models. Rate of Application Value Unit Delayed-release supplement 0.05 % of kg substrate Substrate Rate 16.7 kg/m2 Perlite Rate 0.042 m3/m2 Casing Rate 0.038 m3/m2 Watering Rate 0.0013 m3/m2 Harvester Rate 0.1 labor/m2 Labor Shifts 8 hrs/day Harvesting Days 26 days Air Exchange in Growing 1 Growing room/day Room

10.4.1. Cost Model for Satellite Mushroom Production System

The cost model for the SMPS begins by providing an output for the growing room size required to produce 20 000 kg of mushrooms given a yield of 14.28 kg/m2. Next, the

mass of individual substrate components (commercial grain spawn and supplement) are

calculated. Tray requirements are presented, i.e. amount of perlite needed to cover the

bottom of the trays for the entire growing area, peat, lime, and activated carbon to form

the casing layer. The amount of steam to pasteurize the casing material is calculated by

dividing the heat required to increase the temperature of the casing to 70 oC by the change of enthalpy for steam at 100 oC to 45 oC. The amount of heat lost to the environment for the duration of the 6 hrs pasteurization period is incorporated into the total amount of steam required. The steam cost ($/kg) is calculated based on the cost of

184 generation from natural gas. The cost of the environmental control to cool the growing

room temperature to 18 oC with an inlet air temperature of 25 oC and a heat generation

constant of 0.05 kW/kg substrate is calculated. Table 10.4 provides the values for heating

and energy generation component of the model.

Table 10.4 Parameters used for the energy calculations of the SMPS. Rate of Application Value Unit Water Specific Heat 4.2 kJ/kgoK Peat Specific Heat 3.6 kJ/kgoK Initial Temp. Peat 24 oC Initial Water Temp. 15 oC Pasteurizing Temp. 70 oC Steam Temp. for 100 oC Pasteurization Enthalpy of Steam at 100 oC 2676 kJ/kg Natural Gas Price 12 $/MMBtu Energy Content of Natural 1,055,055 kJ Gas Air Ambient Temp. 25 oC Growing Room Temp. 18 oC Air Specific Heat 1.012 kJ/kgoC Biomass Heat Generation 0.05 kW/kg Substrate day

Finally, the direct costs of the substrate components, casing, perlite, labor, steam,

water, and electricity are calculated. The total direct costs based on these inputs are

$79,795. Mushroom producers would lose $37,195 if they sell mushrooms at the current price of $2.13/ kg mushrooms. The breakdown in costs as a percentage of total direct

costs is depicted in Figure 10.7.

185 Other 0%

Labor 28%

Subs. Req. Tray Req. Labor Other Tray Req. 6% Subs. Req. 66%

Figure 10.6 Breakdown of costs in % of total costs for the SMPS for the substrate requirements, tray requirements, labor and other costs (steam, electricity, and water)

A sensitivity analysis was made for yield (kg/m2) and cost of grain spawn. If mushroom yield is increased to 25-35 kg/m2, a substantial drop in mushroom production costs is encountered. Figure 10.8 depicts the change of mushroom production cost when yield per unit area is increased.

186 12

10

8

($/kg) 6 Mushroom Cost 4

2

0 5 101520253035 Yield (kg/m2)

Figure 10.7. Change of mushroom production cost ($/kg) when yield per unit area (kg/m2) is increased for the SMPS.

The sensitivity analysis shows an almost linear increase in the price of mushrooms when the price of commercial grain spawn is increased. Since the commercial grain spawn is such a large component of the substrate, it is reasonable to assume that this is an important factor that would determine the price of mushrooms in

$/kg. Figure 10.9 depicts the change in mushroom cost ($/kg) as the price of commercial grain spawn ($/kg) is increased.

187 4.5

4

3.5

3

2.5

($/kg) 2 Mushroom Cost 1.5

1

0.5

0 0.2 0.5 0.7 0.9 1.2 1.4 1.6 1.7 1.9 2.1 2.3

Commercial Grain Spawn Cost ($/kg) Figure 10.8. Change in mushroom price ($/kg) with respect to the change in price for commercial grain spawn ($/kg) for the SMPS.

10.4.2. Cost Model for Complete On-site Mushroom Production System

As in the SMPS cost model, the model for the COMPS starts with calculating the characteristics of the growing room. The total components of the grain/oilseed substrate are calculated based on the specified formulation that was used for the control treatments in Scytalidium thermophilum incubation studies. The difference between the cost model for the SMPS and the COMPS is that the former includes the energy, water, and other materials necessary to process the grains onsite. Therefore, all the energy and water requirements are calculated for the different stages of the aseptic processing unit. The first stage of the aseptic processing unit is where the steeping of the grains occurs. The

188 grains are added with the appropriate volume of make-up water at 70 oC needed to increase the moisture content of the grains to 0.6 (wb). Then the amount of steam required to sterilize the substrate was calculated based on a processing temperature of

121 oC. The amount of water needed to cool the grain substrate in the cooling section was

calculated. Steam cost was calculated based on energy generation from natural gas. In

addition, the energy needed to maintain a growing room temperature of 18 oC with an

assumed ambient (inlet) temperature of 25 oC and biological heat output of 0.05-kW/kg

substrate day was calculated. Table 10.5 provides the summary for the energy

calculations used for this cost model.

Table 10.5. Parameters used for the energy calculations of the COMPS. Rate of Application Value Unit Water Specific Heat 4.2 kJ/kgoK Peat Specific Heat 3.6 kJ/kgoK Initial Temp. Peat 24 oC Initial Water Temp. 15 oC Pasteurizing Temp. 70 oC Steam Temp. for 100 oC Pasteurization Enthalpy of Steam at 100 oC 2676 kJ/kg Grain Sterilization Temp. 121 oC Steam Enthalpy at 121 oC 2706 kJ/kg Natural Gas Price 12 $/MMBtu Energy Content of Natural 1,055,055 kJ Gas Air Ambient Temp. 25 oC Growing Room Temp. 18 oC Air Specific Heat 1.012 kJ/kgoC Biomass Heat Generation 0.05 kW/kg Substrate day Specific Heat Grain 2.63 kJ/kgoC Specific Heat Steeped Grain 3.44 kJ/kgoC Specific Heat Soybean 2.14 kJ/kgoC Specific Heat Mineral 2.14 kJ/kgoC

189 Finally, the direct costs of substrate components, labor, steam, electricity, and water are calculated and presented. Total direct costs for the COMPS are $27,908 with a total profit of $14,691. Figure 10.10 depicts the breakdown in costs for the COMPS.

Other 1%

Subs. Req. 37% Subs. Req. Tray Req. Labor Labor 52% Other

Tray Req. 10%

Figure 10.9. Breakdown of costs in % of total costs for the COMPS for the substrate requirements, tray requirements, labor and other costs (steam, electricity, and water)

A sensitivity analysis was performed by varying mushroom yield (kg/m2) in order to observe the effect on mushroom production cost ($/kg). Varying yield from 10-

20 kg/m2 tremendously reduced the cost of mushrooms. From 20-35 kg/m2, the reduction in cost is not as steep. Figure 10.11 depicts the change in mushroom price ($/kg) with respect to a change in mushroom yield (kg/m2).

190 3.5

3

2.5

2

($/kg) 1.5 Mushroom Cost 1

0.5

0 10 15 20 25 30 35 Yield (kg/m2)

Figure 10.10. Change of mushroom production cost ($/kg) when yield per unit area (kg/m2) is increased for the COMPS.

A second sensitivity analysis was done by varying the cost of millet, and it was

found that mushroom production cost varied linearly with the change in cost for the millet grain. Figure 10.12 depicts the change in mushroom production costs with a change in the price of millet grains.

191 1.8 1.6 1.4 1.2 1

($/kg) 0.8

Mushroom Cost 0.6 0.4 0.2 0 0.2 0.4 0.6 0.8 1 1.2 1.4 1.6

Yield (kg/m2)

Figure 10.11. Change in mushroom price ($/kg) with respect to the change in price for millet grains ($/kg) for the COMPS.

10.5. Discussion

Adopting a non-composted grain-based mushroom production system would eventually eliminate the need for the malodorous and lengthy composting process in mushroom production. Furthermore, the resulting waste-stream, i.e. spent grain substrate, could be sold, generating additional income to mushroom producers. The spent grain substrate may be valuable either as an animal feed or as a bio-energy feedstock.

Studies have shown that lingnocellulosic-based spent mushroom substrate from shiitake and oyster mushroom production could potentially be used for animal feed by increasing nutritive values and substrate digestibility (Bisaria et al., 1997; Royse and

Sanchez, 2007). A study by Torev (1968) explored the potential of using Agaricus bisporus mycelium as animal feed. Torev indicated that mushroom mycelium contains about 45% protein. When used as a bio-concentrate ranging from 3%-5% in pig, calf, and

192 broiler feed, an 18% to 22% increase in growth was detected. For laying hens, including

mushroom mycelium in their feed increased egg production 14% to 18%. Hence, the gain

substrate waste-stream could potentially generate additional income for mushroom producers.

The main advantage of the SMPS over the COMPS is that an on-site aseptic processing system is not needed. Hence, the capital investment and operating costs of the aseptic processing system are eliminated for the grower. Furthermore, the technical know-how to operate the aseptic processing system is not needed. The majority of the processing is handled upstream by mushroom spawn producers. Furthermore, this work has shown that older commercial grain spawn has lower productivity than freshly manufactured grain spawn. Hence, it is important to utilize the grain substrate in a timely manner. Transportation of grain spawn material is also an important factor as climate- controlled or refrigerated transportation systems are necessary to prevent vegetative overgrowth and stress on the mushroom fungus. Additionally, mushroom growers using mushroom spawn substrate may be limited to specific proprietary formulations offered by spawn producers with little or no choice of ingredients. However, the most important issue with this system is that spawn producers can charge a premium for the commercial grain spawn they supply and this is why the SMPS is expensive to operate at current grain spawn prices.

The major advantage of adopting the COMPS is the freedom of choosing the specific grain formulations, reduced transportation costs, and freshly produced substrate.

Furthermore, this system will give mushroom producers increased opportunity for growing a variety of fungal organisms used in the nutraceutical, biocontrol, or

193 pharmaceutical industry. Hence, the mushroom farm can become a major supplier of fungal bio-products. For example, producing human therapeutic proteins using a mushroom platform based on solid-state fermentation could yield an economical alternative to liquid fermentation. However, as mentioned earlier, the COMPS has higher capital cost and the technical know-how to operate and maintain such a system could be substantial. Overall, costs of operation are much lower than for the SMPS, and mushroom producers are capable of making a profit if this system is adopted.

10.6. Conclusions

A theoretical design for two mushroom production systems using non-composted grain-based substrates as alternatives to the traditional mushroom production process is presented herein. The SMPS uses commercial grain spawn substrates procured from spawn manufacturers. The major on-site activity in such a system is strictly focused beyond the substrate preparation phase. The COMPS entails the on-site processing and colonization of the grain, and mushroom production.

Based on the cost model developed herein. the cost of the first system is too high to operate at current input values and would lead to losses. However, the COMPS holds the promise to be economically sustainable, and adopting it would eventually eliminate the environmental impact of current conventional compost-based mushroom production.

194 Chapter 11

Conclusions and Scope for Future Research

In this research, an A. bisporus mushroom production system, based on non-

composted grain-based substrates composed of a mixture of commercial grain spawn or

grain/oilseeds supplemented with delayed-release nutrient supplements, was developed as

a potential replacement to the environmentally problematic compost-based system. It was

also shown that a grain-based system could be adapted for the production of other mushroom producing fungi.

In the initial stages of the research, various factors were tested to see if they contributed significantly to an increase in mushroom yield for commercial grain spawn substrates. The first study showed that adding delayed-release nutrient supplements

increased mushroom yield. There was clear indication that out of the two supplements

used (S41 and S44), mushroom yield from S41 was higher. Furthermore, including a

layer of water-holding materials beneath the grain-based substrate increased mushroom

yield from 7.5 kg/m2 to 13 kg/m2. Sterilizing the casing and adding activated carbon (AC)

also significant increased mushroom yield.

In the second study, it was observed that the yield of A. bisporus mushrooms was

not significantly affected by the type of water-holding material placed beneath the substrate. However, on average, perlite produced the highest mushroom yield (8.68 kg/m2) and was used in all additional testing. S41 rate, perlite volume, and casing type

(sterilized or non-sterilized with an AC amendment) were evaluated for their effect on

mushroom productivity. Overall, all three factors had a significant effect on the yield of

195 mushrooms (p<0.05). On average, yields of treatments with non-sterilized casing were

lower than with sterilized casing with AC.

In the third study, adding delayed-release supplements to a basal substrate

consisting of commercial grain spawn increased mushroom yields compared to a

substrate composed solely of grain spawn. Rate and type of supplement were significant.

Among the five delayed-release supplements tested, S41 produced the highest yield and

substrate bioefficiency. The 20% S41 produced 13.73 kg/m2, with a substrate

bioefficiency of 133.7%. Overall, increasing the rate of supplementation above 5% did

not significantly increase yield.

The fourth study showed that the effect of adding AC to heat-treated casing on A. bisporus mushroom production for compost-based substrates was different when

compared to grain spawn-based substrates. For composted substrates that are not fully

colonized with A. bisporus, the addition of AC had no significant effect on mushroom

yield or size (p>0.05). However, casing heat treatment time did increase mushroom yield.

The yield of mushrooms from grain spawn substrates also was sensitive to heat treatment

duration and, in contrast to compost-based substrate, benefited from the addition of AC to

the casing. The highest yield was found for casing containing 5-10% AC. The highest average yield among all treatments with commercial grain spawn was 14.28 kg/m2. A

corresponding BE of 177% was observed for the 5% S41 + 2000 ml perlite + sterilized

casing with activated treatment.

In the fifth study, Agaricus bisporus and A. blazei mushrooms were successfully produced on sterilized substrates consisting of millet grains and three types of oilseeds

(soybean, niger, and safflower). Overall, mushroom yield was influenced by the type and

196 rate of oilseed mixed with millet grain prior to colonization by the mushroom fungus and

by the further addition of S41 after colonization of the substrate. The highest average

yield of mushrooms for a 2-2.5 cm depth of grain/oilseed substrate for A. bisporus was

16.9 kg/m2. The highest yield obtained for a grain/oilseed substrate with A. blazei was

15.9 kg/m2, which compares favorably with reported yields in the range of 17 kg/m2 on composted substrates

In the sixth study, it was observed that mushroom yield, substrate bioefficiency, and spawn-run duration in non-composted grain-based substrates were enhanced to varying degrees by the addition of S. themophilum. Oats, millet, and rye grain substrate

were all shown to support high mushroom yields (17.9, 21.3, and 19.4 kg/m2,

respectively). One important observation is the decrease in spawn-run durations for

treatments with S. thermophilum, compared to controls without S. thermophilum which

ultimately shortens the production cycle. The effect of S. thermophilum in these different

grain substrates was dependent on the type of grain used. Oat-based substrates benefited

the most from the addition of S. thermophilum, i.e. mushroom yield and bioefficiency

increased, whereas rye and millet-based substrates did not. Furthermore, incubation with

S. thermophilum for a period greater than 10 days decreased mushroom yield.

In the seventh study, moisture content of rye-grain based mushroom substrates

was shown to have a direct effect on oxygen uptake rate (OUR), hyphal extension rate,

mushroom yield, and substrate bioefficiency. OURs and hyphal extension rates increased with increasing moisture contents, with highest OUR of 4.88 mg O2/g VS day observed

for the 65% moisture treatment. OUR measurements using pressure sensor methods

appears to negatively impact the growth of the mushroom fungus and it is recommended

197 that continuous respirometric measurement methods are used to ensure that aerobic conditions are maintained throughout the test duration, and that the stimulating effect of carbon dioxide on fungal biomass growth is maintained at optimal levels. Mushroom yield and substrate bioefficiency were highest for substrate moisture contents ranging from 50-65%, whereas 45% and 70% moisture contents were not suitable.

Finally, two theoretical designs for grain-based mushroom production systems were developed, referred to as “Complete On-site” and “Satellite” Mushroom Production

Systems. The first mushroom production system is adapted for the grain/oilseed substrate

and entails the use of an aseptic processing unit. The second is adapted for the

commercial grain spawn substrate that is procured from suppliers of mushroom grain

spawn. Based on the cost model that includes operational costs only, the Complete On- site Mushroom Production Process can provide economically viable yields of mushrooms, and adopting it would provide profits to mushroom producers as opposed to the Satellite Mushroom Production Process. Incorporating capital costs in the cost model may result in a different outcome.

More work is needed in a variety of areas that were not covered by the studies reported herein. One area is the further refining of substrate formulations to increase mushroom yield. It is recommended that statistical-based optimization procedures are used in the substrate formulation process. Furthermore, another area that could be pursued is to determine if there are any interaction effects between the substrate formulation and the casing layer. It was shown the addition of activated carbon to heat- treated casing is required for the commercial grain spawn substrate. However, this may not be the case for the grain/oilseed substrate. This is important because activated carbon

198 is an expensive additive to the casing and eliminating its use will decrease costs.

Work with S. thermophilum has shown that its addition benefits the growth of A.

bisporus. However, results were inconclusive on whether S. thermophile can confer

selectivity to A. bisporus growth in grain-based substrates. However, there is a multitude

of thermophiles that inhabit mushroom compost that may provide this selectivity.

Furthermore, there are a variety of fungal biocontrol agents that could be tested to see whether mushroom pathogens can be inactivated while maintaining adequate mushroom growth. Ultimately, this would help in eliminating the need for sterilizing the grain-based substrate; further reducing costs of the process.

Another important area that needs further work is the development of a procedure to process the grains and model the different steps in the proposed theoretical aseptic processing system that is based on segmented-flow technology. In addition, it would be important to do more work in the substrate flowability measurements since this is a crucial parameter that determines whether a substrate is easy to handle or not.

Finally, a pilot scale system should be developed since most tests until now have been confined to a lab-scale size. Ultimately, this would be the last step before moving the grain-based system to a commercial-scale level if this ever were to come to fruition.

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219

Appendix A

Cultivation of Pleurotus eryngii on Substrates Composed of Grains and Oilseeds

Introduction

Pleurotus eryngii, the King Oyster mushroom, production is steadily increasing

with production levels in Japan up to 29,882 tons, whereas production in the United

States for 2004 was estimated to be 85,000 kg (Yamanaka, 2005; Royse et al., 2005). The

US market price for P. eryngii was estimated at $9-11/ kg fresh (Royse et al., 2005). Tan

et al. (2005) describes three methods of cultivation: bag, casing, and bottle. The bottle

method, is recommended for large scale operations because it can be mechanized.

Substrates used for the cultivation of the P. eryngii are composed of mixtures of wheat

straw, sawdust, cotton seed hulls supplemented with bran, corn powder, sliced sugar

beets or fruits and gypsum (Rodriguez, 2005). The substrate is either pasteurized or

sterilized as in the case of the bottle system. Spawn runs in the bottle system last about 35

days at 22-23 °C during which the mushroom fungus colonizes the substrate. The highest

observed yield by Rodriguez and Royse (2006) was 135 g for a substrate composed of

cottonseed hulls (62%), corn distiller’s grain (4%), calcium sulphate (1%), ground

soybean (6%), aged red oak sawdust (27%), and supplemented with 50 μg/g substrate of

manganese.

In this study, grain-based substrates supplemented with oilseeds were tested as a

substrate for the production of P. eryngii.

220

Methods

Microorganism and Inoculum Preparation

Pleurotus eryngii (MC888) pure culture grown on agar was obtained from the

Mushroom Spawn Lab (PSU). As inoculum rye grain spawn (200 g), calcium carbonate

(4 g), calcium sulfate (4 g), and water (220 ml) were added to 1000 ml wide mouth

Erlenmeyer flasks. The flasks were sealed with cottonwool plugs and autoclaved for 60

min. The flasks were transferred to a laminar flow hood and allowed to cool overnight.

Four agar plugs were aseptically added to each flask and then the flasks were placed at 23

°C. The flasks were periodically shaken once a week to completely to hasten

colonization. On average it took 14 days for the grains to be fully colonized after which

the material was stored at 5 °C until use.

Bottle Preparation Process

Grains and oilseeds were placed in polypropylene (L = 0.37 m x W = 0.25 m x D

= 0.14 m) containers with the appropriate amount of water in an autoclave for 45 min to

achieve a final moisture content of 0.6 (wet basis). The cooked grains (500 g) along with

the appropriate amount of oilseed (5% of the substrate formulation- 50 g), and calcium

sulfate (25 g) were mixed and then placed in 500 ml Ball® Jars. The materials were then

sealed using Tyvac® and attached in place using the screw top of the jar. The jars were

then autoclaved for 2 hrs. Following, the jars were placed in a laminar flow hood and

allowed to cool. The jars were inoculated with 10 g of rye grain spawn and then placed in

a mushroom production chamber.

221 Environmental Conditions in Mushroom Production Chamber

The inoculated bottles were set in a mushroom production chamber were the temperature was controlled at 22 °C. Once the substrate was fully colonized, the bottles were transferred to an indoor greenhouse (Midwest Quality Qlove Inc, Chillicothe, MO) covered with polyethylene in which ambient temperature was maintained at 20 °C using an air conditioning system. Carbon dioxide was maintained below 800 ppm, and humidity was supplied by a humidifier and ranged from 80-90%. Humidification was controlled using an on-off timer to achieve a 12hr/day humidification cycle. A light source (100 W) was added over the greenhouse and maintained 24 hrs/day. Aeration was achieved using an air pump with an aeration rate of 10 l/min/24 hrs. Carbon dioxide, relative humidity, and temperature were monitored using the following instruments

Extech, model 444712, Extech Instruments, Waltham, Mass) for temperature and humidity and Telair 7001 (Goleta, CA 93117) for carbon dioxide.

Experimental Set-up

The first set of experiment involved a factorial design testing substrates composed of different cereal grains pre-treated to contain 60% moisture content supplemented with different wetted oilseeds at 5% of the substrate total composition. In some treatments the oilseeds were blended using a commercial blender to form smaller particles. The grains that were tested were the following: oats, rye, and millet. The oilseeds tested were: cracked roasted soybean, sunflower, and safflower.

Results and Discussion

The overall results indicate that production of P. eryngii on grain-based substrates is possible. Rye grain did not support adequate mushroom yield in most treatments. This

222 is not surprising since P. eryngii is typically grown on sawdust or cottonseed hull based

substrate that are rich in cellulose. Since P. eryngii is a primary decomposer (white rot

fungus) the availability of cellulose in the substrate is important. Therefore, a starch-

based substrate seems to be ineffective in supporting adequate mushroom yields. In

contrast, oat-based substrate which are rich in fiber (cellulose) content supported

adequate mushroom yield with the highest yield observed for the control group (no

oilseed addition) of 106 g. Table 1 summarizes the results for the different grain-based

substrates supplemented with 3 oilseeds, whereas Figure 1 depicts an image of P. eryngii

fruiting on oat-based substrates.

Table A1. Summary of Pleurotus eryngii yield from grain-based substrates supplemented with oilseed. Cereal Grain Oilseed Mushroom Yield (g) Oat - 106 ±11 Soybean 86 ± 36 Soybean-blended 93 ±16 Safflower 102 ± 9 Safflower-blended 99 ± 3 Sunflower 47 ± 23 Sunflower-blended 76 ± 21 Rye - 42 ± 7 Soybean 0 ± 0 Soybean- blended 0 ± 0 Safflower 0 ± 0 Safflower-blended 0 ± 0 Sunflower 53 ± 47 Sunflower-blended 19 ± 20 Millet - 54 ± 16 Soybean 63 ± 13 Soybean- blended 86 ± 13 Safflower 59 ± 18 Safflower-blended 73 ± 24 Sunflower 67 ± 27 Sunflower-blended 60 ± 16.5

223

Figure A1. Pleurotus eryngii fruiting on oat-based substrates

Conclusions

P. eryngii was shown to fruit on grain-based substrates, and this provides evidence that a variety of mushroom can be grown on substrate primarily composed of

grains. Oat-based substrates appear to best suite P. eryngii mushroom production and

highest yield of mushrooms was 106 g. More work is needed to further improve yield of

P. eryngii in grain-based substrates.

References

Rodriguez-Estrada, A. E. 2005. Influence of substrate composition and mushroom strains

on productivity and susceptibility of Pleurotus eryngii to bacterial blotch disease.

224 MS thesis. University Park., PA. The Pennsylvania State University, Department

of Plant Pathology.

Rodriguez-Esrada, A. E., and D. J. Royse. 2006. Yield, size, and bacterial blotch

resitance of Pleurotus eryngii grown on cottonseed hulls/oak sawdust

supplemented with manganese copper and whole ground soybean. Biores. Tech.

available at: www.sciencedirect.com. Accessed November 2006.

Royse, D. J., Q. Shen, and C. McGarvey. 2005. Consumption and production of recently

domesticated edible fungi in the United States with a projection of their potential.

In Proc. 5th International Conference on Mushroom Biology and Mushroom

Products, 331-337. Shanghai, China: Acta Edulis Fungi.

Tan, Q., Z. Wang, J. Cheng, Q Guo, L. Guo. 2005. Cultivation of Pleurotus spp. in China.

In Proc. of the Fifth International Mushroom Conference on Mushroom Biology

and Mushroom Products, 338-349. Shanghai, China: Acta Edulis Fungi

Yamanaka, K. 2005. Cultivation of New Mushroom Pleurotus species in East Asia. In

Proc. 5th International Conference on Mushroom Biology and Mushroom

Products, 343-349. Shanghai, China: Acta Edulis Fungi.

225

Appendix B

Rye Grain Substrate Flowabilty

Introduction

One important factor that has been observed but not quantified by mushroom

spawn producers is the change in grain flowability induced by different grain moisture

levels. Flow is defined as the relative movement of a bulk of particles (grains in this case) with respect to neighboring particles or along a container wall surface (Pelleg, 1977).

Flowability can be measured based on the mass flow rate through a conical funnel

(Barbosa and Yan, 2003). In addition, material flowability can be measured using several other parameters such of angel of repose, bulk density, angle of interaction friction,

cohesion, adhesion, and compressibility (Ganesan et al., 2005). A flexible cubical triaxial

tester (CTT) measures fundamental mechanical responses by applying pressures two 6

sides (cubical form) of a given sample (Kamath and Puri, 1997). The failure stress is

what can be obtained from the CTT i.e. material which has small failure stress are more

flowable than material characterized with a large failure stress. This is based upon the

idea that for granular material to flow, the material has to break apart (loss of cohesiveness) for it to flow. Mushroom spawn producers refrain from increasing grain moisture levels beyond 52% because of problems of grain flowability and the formation what is collectively termed as a “stuck reactor”. This increase in stickiness is due to the gelatinization of starch and the leaching of starch granules out of the grains.

226 A flexible-boundary cubical triaxial tester (CTT) developed and described by

Kamath and Puri (1997) was used to measure fundamental mechanical responses of the

grain substrates. In summary, this CTT applies pressure on six sides of a cubical sample

(grain) via flexible rubber membranes using compressed air. The sample holder is cube- shaped with dimensions of 50.8 × 50.8 × 50.8 mm and covered with rubber membranes.

Due to the application of pressure, the sample undergoes changes in dimensions which are recorded using linear motion potentiometers that are in constant contact with pressure application membranes, i.e., all six faces.

Methods

Sample Preparation

Rye grain substrates were prepared as described in Chapter 9. In summary, rye grains, water, calcium carbonate and calcium sulfate were added to 1000 ml Erlenmeyer flasks. The flasks were sealed with cotton wool plugs and autoclaved for 45 min (121oC at 103.4 kPa). The moisture levels of the rye grain substrate were 45, 50, 55, 60, 65, and

70% (wb). The samples were allowed to cool overnight right after they were shaken to distribute the grains.

Sample loading and Testing

The rye grain substrates were individually placed in the sample holder by filling the entire void, and the top part of the CTT was bolted and sealed over the sample. The pressure of all six horizontal membranes was increased until the confining pressure was reached and they were maintained at that pressure. The tests were conducted at confining pressures of 3 and 6 kPa. Next, the pressure of the top and bottom membrane was

227 increased until sample failure occurred. In the case that no clear failure point was

observed, a 15% axial strain was taken as the point of failure (ASTM, 2006).

According to the criterion that shear stress increases with increase in the normal

stress on the failure plane, the Mohr coulomb failure criterion can be expressed

mathematically as:

τ = c + σ tanφ Eq. 10.1

where

τ = shear stress on the failure plane

c = cohesion of the material

σ = normal effective stress on the failure surface

φ = angle of internal friction.

The criterion is shown graphically in Figure 10.6

Figure B1. Mohr Coulomb yield criterion

228 c and τ are the parameters to be determined.

The failure criterion can also be written as

σ −σ σ + σ 1 3 = 1 3 sinφ + c cosφ Eq. 10.2 2 2

Results and Discussion of Flowability Test

The strength of material increased with increase in confining pressure, which is expected. The increase in strength was more prominent at low moisture content (45-

55%). For instance, the strength of the grain sample increased from 16 to 20 kPa for the

45% moisture level. At higher moisture levels (60-70%), a small increase in strength was observed. At the 70% moisture level, the increase in strength was only 1 kPa. With increase in moisture content, the grains became soft and hence the strain was mainly compression of the grain instead of failure of the bulk. Table B1 shows the change in failure strength of the grain substrate with the two confining pressures for the various moisture levels.

Table B1. Failure strength of grain substrate at different confining pressures for the various moisture levels. Failure Strength Failure Strength Moisture Treatment (kPa) (kPa) (%) at 3 kPa at 6kPa 45 16 20 50 15 20 55 15 17 60 10 13 65 10 12 70 9 10

An observed change in material property was detected for the moisture level of 60

% and above. The cohesiveness (c) increased and angle of shear resistance (φ ) decreased which is opposite to the expected trend, which may be due to the softening of the

229 material. Therefore, the material behavior was grouped into two categories. One group was for the moisture levels ranging from 45–55%, and the other for moisture levels ranging from 60-70%.

The results from the CTT were used to determine the parameters of the Mohr coulomb model. For soft material (which deforms with the application of pressure), the

Mohr coulomb model cannot explain well the bulk behavior. For moisture contents of

45–55%, the c value increased from 3.95 to 5.02. The increase in grain substrate cohesiveness is an indication that the materials become less flowable. The angle of shear resistance (φ ) decreased with moisture content. For moisture contents of 60-70%, the value increased from 2.48 to 3.46. This also indicates that the grain became less flowable with an increase in moisture, but Mohr coulomb alone cannot be suitable to explain the behavior due to reason stated above. Table B2 provides a summary of the parameters of the Mohr coulomb model.

Table B2. Summary of parameters of the mohr coulomb model Moisture Treatment (%) Cohesiveness φ C-value 45 3.95 23.57 50 4.96 19.46 55 5.02 14.47 60 2.48 19.47 65 3.09 14.47 70 3.46 8.21

References

ASTM. 2006. D4767-04 Standard test method for consolidated undrained triaxial

compression test for cohesive soils. Annual Book of ASTM Standards, American

Society for Testing and Materials.

230 Barbosa-Canovas, G. V., and H. Yan. 2003. Powder characteristics of preprocessed

cereal flours. In Characterization of cereals and flours, 173-209. New York,

N.Y.: Marcel Dekker.

Ganesan, V., K. A. Rosentrater, and K. Muthukumarappan. 2005. Flowability and

handling characteristic of bulk solids and powders. ASABE Paper No. 056023 St.

Joseph, Mich.: ASAE.

Kamath, S., and V. M. Puri. 1997. Measurement of powder flow consecutive model

parameters using a cubicle triaxial tester. Powder Technology 90: 59-70.

Peleg, M. 1977. Flowability of food powders and methods for its evaluation- A review.

Journal of Food Process Engineering 1: 303-328.

231 Appendix C

Cost Model Excel Spreadsheets

232

233

234

235

236

Vita Mark A. Bechara

EDUCATION Pennsylvania State University, University Park PA • PhD in Agricultural and Biological Engineering- Emphasis in Biological/Bioprocess Engineering (12/07). • MS in Agricultural and Biological Engineering- Emphasis in Biological/Bioprocess Engineering (8/04). American University of Beirut, Ras Beirut Lebanon • BS in Agricultural Sciences and Diploma of Ingenieur Agricole (7/01) SCHOLARSHIPS AND AWARDS • First place winner at the 21st Graduate Poster Exhibition 2006. The Pennsylvania State University for: “of Agaricus blazei: A novel mushroom with anti-tumor properties” • Second place winner at the 20th Graduate Poster Exhibition 2005. The Pennsylvania State University for: “Production of Agaricus bisporous on a non- composted substrate”. • Second place winner at the College of Agricultural Sciences and Gamma Sigma Delta research expo 2005, at the Pennsylvania State University for “Production of Agaricus bisporus on a non-composted substrate”. • Member of Alpha Epsilon “The Honor Society for Agriculture Engineering”. • Awarded the “Harold V. and Velma B. Walton Doctoral Student Endowment in Agricultural and Biological Engineering” • Member of the Honorary Society of Agriculture Gamma Sigma Delta. • Third place winner at the 19th Graduate Poster Exhibition 2004, at The Pennsylvania State University for: “Cultivation Systems for the Button Mushroom Agaricus bisporus”. files PUBLICATIONS • Bechara, M. A., P. Heinemann, P. N. Walker, and C. P. Romaine. 2006. Non- composted grain-based substrates for the production of Agaricus bisporus mushroom. Transactions of the ASABE 49(3): 819-824. • Bechara, M. A., P. Heinemann, P. N. Walker, and C. P. Romaine. 2006. Agaricus bisporus cultivation in hydroponic systems. Transaction of the ASABE 49(3): 825-832. • Bechara, M. A., P. Heinemann, C. P. Romaine, P. N. Walker and V. Wilkinson. 2006. Evaluating Non-composted Grain Substrates for the Production of Agaricus bisporus and Agaricus blazei Mushrooms. ASABE Annual International Meeting. Portland, Oregon, USA, July 9-12. • Bechara, M. A., P. Heinemann, P. N. Walker and C. P. Romaine. 2005. Cultivation of Agaricus bisporus on a mixture of cereal grain spawn and delayed- release nutrient supplement. Mushroom News, August 2005.