ECTOMYCORRHIZAL FUNGAL COMMUNITIES ASSOCIATED WITH HIMALAYAN CEDAR FROM PAKISTAN

A dissertation submitted to the University of the Punjab in partial fulfillment of the requirements for the degree of

DOCTOR OF PHILOSOPHY

in BOTANY by

SANA JABEEN

DEPARTMENT OF BOTANY UNIVERSITY OF THE PUNJAB LAHORE, PAKISTAN

JUNE 2016

TABLE OF CONTENTS

CONTENTS PAGE NO. Summary i Dedication iii Acknowledgements iv CHAPTER 1 Introduction 1 CHAPTER 2 Literature review 5 Aims and objectives 11 CHAPTER 3 Materials and methods 12 3.1. Sampling site description 12 3.2. Sampling strategy 14 3.3. Sampling of sporocarps 14 3.4. Sampling and preservation of fruit bodies 14 3.5. Morphological studies of fruit bodies 14 3.6. Sampling of morphotypes 15 3.7. Soil sampling and analysis 15 3.8. Cleaning, morphotyping and storage of ectomycorrhizae 15 3.9. Morphological studies of ectomycorrhizae 16 3.10. Molecular studies 16 3.10.1. DNA extraction 16 3.10.2. Polymerase chain reaction (PCR) 17 3.10.3. Sequence assembly and data mining 18 3.10.4. Multiple alignments and phylogenetic analysis 18 3.11. Climatic data collection 19 3.12. Statistical analysis 19 CHAPTER 4 Results 22 4.1. Characterization of above ground ectomycorrhizal fungi 22 4.2. Identification of ectomycorrhizal host 184 4.3. Characterization of non ectomycorrhizal fruit bodies 186 4.4. Characterization of saprobic fungi found from fruit bodies 188 4.5. Characterization of below ground ectomycorrhizal fungi 189 4.6. Characterization of below ground non ectomycorrhizal fungi 193 4.7. Identification of host taxa from ectomycorrhizal morphotypes 195 4.8. Soil analysis 198 4.9. Community analysis 199 4.9.1. Alpha diversity 199 4.9.2. Beta diversity 205 4.9.3. Gamma diversity 206

CHAPTER 5 Discussion 234 References 242 Annexure

LIST OF FIGURES TITLE PAGE NO. Figure 1. Map of Pakistan showing sampling sites 13 Figure 2. Morphology of ahmadii 26 Figure 3. Anatomy of Amanita ahmadii 27 Figure 4. Molecular phylogenetic analysis of Amanita ahmadii based on ITS sequences 28 Figure 5. Molecular phylogenetic analysis of Amanita ahmadii based on LSU sequences 29 Figure 6. Morphology of Amanita brunneopantherina 32 Figure 7. Anatomy of Amanita brunneopantherina 33 Figure 8. Molecular phylogenetic analysis of Amanita brunneopantherina based on ITS sequences 34 Figure 9. Morphology of Amanita flavipes 37 Figure 10. Anatomy of Amanita flavipes 38 Figure 11. Molecular phylogenetic analysis of Amanita flavipes based on ITS sequences 39 Figure 12. Morphology and anatomy of Amanita glarea 42 Figure 13. Molecular phylogenetic analysis of Amanita glarea based on ITS sequences 43 Figure 14. Molecular phylogenetic analysis of Amanita glarea based on LSU sequences 44 Figure 15. Morphology of Amanita swatica 47 Figure 16. Anatomy of Amanita swatica 48 Figure 17. Molecular phylogenetic analysis of Amanita swatica based on ITS sequences 49 Figure 18. Morphology of Boletus himalayensis 52 Figure 19. Anatomy of Boletus himalayensis 53 Figure 20. Molecular phylogenetic analysis of Boletus himalayensis based on ITS sequences 54 Figure 21. Molecular phylogenetic analysis of Boletus himalayensis based on LSU sequences 55 Figure 22. Morphology and anatomy of rubellus 58 Figure 23. Molecular phylogenetic analysis of Hortiboletus rubellus based on ITS sequences 59 Figure 24. Morphology and anatomy of luridiformis 62 Figure 25. Molecular phylogenetic analysis of Neoboletus luridiformis based on ITS sequences 63 Figure 26. Morphology of rimosus 66 Figure 27. Anatomy of Xerocomellus rimosus 67 Figure 28. Molecular phylogenetic analysis of Xerocomellus rimosus based on ITS sequences 68 Figure 29. Molecular phylogenetic analysis of Xerocomellus rimosus based on LSU sequences 69 Figure 30. Morphology of corrosus 72

Figure 31. Anatomy of Cortinarius corrosus 73 Figure 32. Molecular phylogenetic analysis of Cortinarius corrosus based on ITS sequences 74 Figure 33. Morphology of Cortinarius longistipus 77 Figure 34. Anatomy of Cortinarius longistipus 78 Figure 35. Molecular phylogenetic analysis of Cortinarius longistipus based on ITS sequences 79 Figure 36. Morphology of galiyensis 82 Figure 37. Anatomy of Geastrum galiyensis 83 Figure 38. Molecular phylogenetic analysis of Geastrum galiyensis based on ITS sequences 84 Figure 39. Morphology of flavostipus 87 Figure 40. Anatomy of Gomphidius flavostipus 88 Figure 41. Molecular phylogenetic analysis of Gomphidius flavostipus based on ITS sequences 89 Figure 42. Molecular phylogenetic analysis of Gomphidius flavostipus based on LSU sequences 90 Figure 43. Morphology of Hebeloma angustisporium 93 Figure 44. Anatomy of Hebeloma angustisporium 94 Figure 45. Molecular phylogenetic analysis of Hebeloma angustisporium. based on ITS sequences 95 Figure 46. Morphology of alba 98 Figure 47. Anatomy of Inocybe alba 99 Figure 48. Morphology of Inocybe flavellorimosa 102 Figure 49. Anatomy of Inocybe flavellorimosa 103 Figure 50. Molecular phylogenetic analysis of Inocybe spp. based on ITS sequences 104 Figure 51. Morphology of Inocybe mimica 107 Figure 52. Anatomy of Inocybe mimica 108 Figure 53. Molecular phylogenetic analysis of Inocybe mimica based on ITS sequences 109 Figure 54. Morphology of Inocybe oblectabilis 112 Figure 55. Anatomy of Inocybe oblectabilis 113 Figure 56. Molecular phylogenetic analysis of Inocybe oblectabilis based on ITS sequences 114 Figure 57. Morphology and anatomy of Rhizopogon flavus 117 Figure 58. Molecular phylogenetic analysis of Rhizopogon flavus based on ITS sequences 118 Figure 59. Morphology of amythistina 122 Figure 60. Anatomy of Russula amythystina 123 Figure 61. of Russula amythestina 124

Figure 62. Molecular phylogenetic analysis of based on ITS sequences 125 Figure 63. Morphology of Russula delica 128 Figure 64. Anatomy of Russula delica 129 Figure 65. Molecular phylogenetic analysis of Russula delica based on ITS sequences 130 Figure 66. Morphology of Russula pakistanica 136 Figure 67. Scanning electron micrographs of of Russula pakistanica 137 Figure 68. Anatomy of Russula pakistanica 138 Figure 69. Ectomycorrhiza of Russula pakistanica 139 Figure 70. Molecular phylogenetic analysis of Russula pakistanica based on ITS sequences 140 Figure 71. Molecular phylogenetic analysis of Russula pakistanica based on LSU sequences 141 Figure 72. Morphology of Russula rubecola 144 Figure 73. Anatomy of Russula rubecola 145 Figure 74. Molecular phylogenetic analysis of Russula rubecola based on ITS sequences 146 Figure 75. Morphology of convexatus 149 Figure 76. Anatomy of Suillus convexatus 150 Figure 77. Molecular phylogenetic analysis of Suillus convexatus based on ITS sequences151 Figure 78. Morphology of terreum 154 Figure 79. Anatomy of 155 Figure 80. Molecular phylogenetic analysis of Tricholoma terreum based on ITS sequences 156 Figure. 81. Morphology of khanspurensis 159 Figure 82. Anatomy of Gyromitra khanspurensis 160 Figure 83. Molecular phylogenetic analysis of Gyromitra khanspurensis based on ITS sequences 161 Figure. 84. Morphology of pakistanica 164 Figure 85. Anatomy of Morchella pakistanica 165 Figure 86. Molecular phylogenetic analysis of Morchella pakistanica based on ITS sequences 166 Figure. 87. Morphology of asiatica 169 Figure 88. Anatomy of Verpa asiatica 170 Figure 89. Molecular phylogenetic analysis of Verpa asiatica based on ITS sequences 171 Figure 90. Molecular phylogenetic analysis of Verpa asiatica based on LSU sequences 172 Figure. 91. Morphology of khanspurensis 175 Figure 92. Anatomy of Peziza khanspurensis 176 Figure 93. Molecular phylogenetic analysis of Peziza khanspurensis based on ITS sequences 177 Figure. 94. Morphology of Geopora pinyonensis 181

Figure 95. Anatomy of Geopora pinyonensis 182 Figure 96. Molecular phylogenetic analysis of Geopora pinyonensis based on ITS sequences 183 Figure 97. Molecular phylogenetic analysis of ectomycorrhizal hosts based on ITS sequences 185 Figure 98. Graph showing absolute abundance of above ground ectomycorrhizal fungal taxa in stand 1 208 Figure 99. Pie chart showing relative abundance of above ground ectomycorrhizal fungal taxa in stand 1 208 Figure 100. Rank abundance curve of above ground ectomycorrhizal fungal taxa in stand 1 209 Figure 101. Graph showing absolute abundance of below ground ectomycorrhizal fungal taxa in stand 1 209 Figure 102. Pie chart showing relative abundance of below ground ectomycorrhizal fungal taxa in stand 1 210 Figure 103. Rank abundance curve of below ground ectomycorrhizal fungal taxa in stand 1 210 Figure 104. Graph showing absolute abundance of above ground ectomycorrhizal fungal taxa in stand 2 211 Figure 105. Pie chart showing relative abundance of above ground ectomycorrhizal fungal taxa in stand 2 211 Figure 106. Rank abundance curve of above ground ectomycorrhizal fungal taxa in stand 2 212 Figure 107. Graph showing absolute abundance of below ground ectomycorrhizal fungal taxa in stand 2 212 Figure 108. Pie chart showing relative abundance of below ground ectomycorrhizal fungal taxa in stand 2 213 Figure 109. Rank abundance curve of above ground ectomycorrhizal fungal taxa in stand 2 213 Figure 110. Graph showing absolute abundance of above ground ectomycorrhizal fungal taxa in stand 3 214 Figure 111. Pie chart showing relative abundance of above ground ectomycorrhizal fungal taxa in stand 3 214 Figure 112. Rank abundance curve of above ground ectomycorrhizal fungal taxa in stand 3 215 Figure 113. Graph showing absolute abundance of below ground ectomycorrhizal fungal taxa in stand 3 215 Figure 114. Pie chart chowing relative abundance of below ground ectomycorrhizal fungal taxa in stand 3 216 Figure 115. Rank abundance curve of below ground ectomycorrhizal fungal taxa in

stand 3 216 Figure 116. Graph showing absolute abundance of above ground ectomycorrhizal fungal taxa in stand 4 217 Figure 117. Pie chart showing relative abundance of above ground ectomycorrhizal fungal taxa in stand 4 217 Figure 118. Rank abundance curve of above ground ectomycorrhizal fungal taxa in stand 4 218 Figure 119. Graph showing absolute abundance of below ground ectomycorrhizal fungal taxa in stand 4 218 Figure 120. Pie chart showing relative abundance of below ground ectomycorrhizal fungal taxa in stand 4 219 Figure 121. Rank abundance curve of below ground ectomycorrhizal fungal taxa in stand 4 219 Figure 122. Graph showing absolute abundance of above ground ectomycorrhizal fungal taxa in stand 5 220 Figure 123. Pie chart showing relative abundance of above ground ectomycorrhizal fungal taxa in stand 5 220 Figure 124. Rank abundance curve of above ground ectomycorrhizal fungal taxa in stand 5 221 Figure 125. Graph showing absolute abundance of below ground ectomycorrhizal fungal taxa in stand 5 221 Figure 126. Pie chart showing relative abundance of below ground ectomycorrhizal fungal taxon in stand 5 222 Figure 127. Rank abundance curve of below ground ectomycorrhizal fungal taxa in stand 5 222 Figure 128. Graph showing absolute abundance of below ground ectomycorrhizal fungal taxa in stand 6 222 Figure 129. Pie chart showing relative abundance of below ground ectomycorrhizal fungal taxa in stand 6 223 Figure 130. Rank abundance curve of below ground ectomycorrhizal fungal taxa in stand 6 223 Figure 131. Box and whisker plot showing the number and distribution of above ground taxa in different stands 224 Figure 132. Box and whisker plot showing the number and distribution of below ground taxa in different stands 224 Figure 133. Above ground accumulation curve for Hazara division 227 Figure 134. Below ground species accumulation curve for Hazara division 227 Figure 135. Above ground species accumulation curve for Malakand division 228 Figure 136. Below ground species accumulation curve for Malakand division 228 Figure 137. Above ground species accumulation curve for Rawalpindi division 229

Figure 138. Below ground species accumulation curve for Rawalpindi division 229 Figure 139. Above ground community composition and comparison of stands based on frequency of taxa 230 Figure 140. Below ground community composition and comparison of stands based on frequency of taxa 231 Figure 141. Above and below ground representative genera of ectomycorrhizal fungal taxa 232 Figure 142: Canonical correspondence analysis of above ground data with edaphic factors 233 Figure 143. Canonical correspondence analysis of below ground data with edaphic factors 233

LIST OF TABLES TITLE PAGE NO. Table 1. List of formulae used for the calculation of diversity indices 21 Table 2. Non ectomycorrhizal fruit bodies collected from different stands 186 Table 3. Saprobic fungi found from fruit bodies 188 Table 4. List of below ground ectomycorrhizal fungi/OTUs 189 Table 5. List of below ground non ectomycorrhizal fungi 193 Table 6. Host tree species identified from ectomycorrhizal morphotypes 195 Table 7. Stand wise results of different soil analysis parameters 198 Table 8. Stand wise above and belowground alpha diversity of the taxa 225 Table 9. Similarity and dissimilarity indices of above ground data 226 Table 10. Similarity and dissimilarity indices of below ground data 226

SUMMARY The present research project deals with the ectomycorrhizal fungal communities associated with Himalayan cedar (Cedrus deodara) from Pakistan. The community study was based on characterization of above and below ground taxa found in association with cedar. Characterization was based on molecular characters using different molecular markers combining with morphological comparisons. Six sampling stands; Khanian forest (stand 1), Khanspur forest (stand 2), Kalam forest (stand 3), Mashkun forest (stand 4), Kuzah Gali (stand 5) and Patriata (stand 6) in Hazara and Malakand division of Khyber Pakhtunkhwa, and Rawalpindi division of Punjab province were selected as sampling sites with distinct climatic conditions. These forests were dominated by C. deodara growing with other members of Pinaceae and deciduous trees. Sampling was carried out in three consecutive years (2012–2014). A total of 107 fungal taxa were identified belonging to 67 genera and 50 families. Among these, 69 species belong to 33 genera and 25 families were ectomycorrhizal. From these 69 species, 33 species were identified as ectomycorrhizal fungal fruit bodies belonging to 19 genera and 14 families and 36 species belonging to 14 genera and 11 families were identified from ectomycorrhizal morphotypes. From the above and below ground data, four species were found as fruit bodies as well as in the form of ectomycorrhizae. So, 65 distinct ectomycorrhizal species were identified and remaining taxa belong to non ectomycorrhizal fungi and excluded in the final dataset for community analysis. The ectomycorrhizal fungal community consisted of a few most frequent fungi and numerous rare taxa. The total species diversity (gamma diversity) was determined by the mean species diversity in each stand (alpha diversity) and the differentiation among these habitats (beta diversity). One way ANOVA was used to analyze the differences among the communities in different stands in major regions from the above and below ground data. The communities did not show significant similarities. From stand 1 and 2, Inocybe was found to be among the most abundant, dominant and diverse . So, in the stands located in Hazara division, Inocybe was found most prevailing genus. From stand 3, Inocybe followed by Russula constituted the major part of the community, and in stand 4, Neoboletus and Amanita were found abundant as fruit bodies while Inocybe and Russula dominated the belowground community. The overall diversity in the stands of Malakand division was represented by 20 genera and 47 species. Among these, Inocybe was the major part of the community

followed by Russula in the form of fruit bodies as well as morphotypes. From stand 5, Inocybe sp. 6 and R. delica were the major community components in this stand. From stand 6, above ground ectomycorrhizal fungal community components were not observed but from the belowground, Russula pakistanica was the most abundant taxon. Thus, the community composition in stands of Rawalpindi division was represented by Inocybe and Russula. It was noted that the host species, age of stand, as well as soil nutrients and organic matter content influenced the distribution patterns of these ectomycorrhizal fungal species. High nutrient availability exhibited lower number of ectomycorrhizal fruit bodies, morphotype abundance, and species richness when compared to nutrient poor areas. Hyper diverse communities were recorded in stand 3 and 4, containing comparatively high phosphorus concentration. Stands with lower electrical conductivity of soil showed a higher level of colonization, species richness and diversity indicated a negative relation. The results also indicated that the rainfall patterns did not affect the fungal communities. The species richness estimators indicated that the level of total species richness was expected to be higher in each stand especially in dryer stands (Rawalpindi division).These results indicated that the forests studied have a great potential for ectomycorrhizal fungal diversity. The ectomycorrhizal symbionts identified during this study could be a source of information for future studies in restoration of rapidly declining forest cover. The data generated during this study could be used for morphological comparisons. Molecular data deposited in different repositories could be used as a reference for future taxonomic and phylogenetic work.

DEDICATED TO My Parents & Teachers Who made my life worth living

ACKNOWLEDGEMENTS All praise is due to Allah, the Almighty, who taught me and gave me strength to know what I knew not. Words are inadequate to express my gratitude to my respected teacher and PhD supervisor, Prof. Dr. Abdul Nasir Khalid, Department of Botany, University of the Punjab, Lahore, Pakistan for his continuous guidance, encouragement and support throughout the duration of my research work. I feel honour to be thankful to Prof. Dr. Muhammad Saleem (Chairman) and Prof. Dr. Khan Rass Masood (Ex-Chairman), Department of Botany, University of the Punjab, Lahore, Pakistan for providing me conducive environment and facilities to carrying out this research project. My sincere thanks to Dr. Abdul Rehman Khan Niazi (Department of Botany, University of the Punjab, Lahore, Pakistan) for his guidance in the field work and helping me in handsome understanding of the subject. I would like to thank Dr. Najam-ul-Sehar Afshan (Center for Undergraduate Studies, University of the Punjab, Lahore), Dr. Muhammad Hanif (Government College University, Lahore), Dr. Tayiba Ashraf (Government Post Graduate College for Women, Sheikhupura), Dr. Samina Sarwar, Dr. Sobia Ilyas (Lahore College for Women University, Lahore), Dr. Abdul Razaq (University of Veterinary and Animal Sciences, Pattoki), and Dr. Syeda Bint-e-Zahira (Government Degree College, Salamatpura, Lahore) for their support and guidance as senior lab members. I am profoundly grateful to Prof. Donald H. Pfister, Department of Organismic and Evolutionary Biology, Harvard University, Cambridge, MA, USA for providing me excellent opportunity to work in his laboratory under his kind supervision. I extend my gratitude to Dr. Katherine Lobuglio, for her guidance in molecular work and valuable discussions to improve the results at Pfister's lab, Harvard University, USA. I am thankful to Dr. Else C. Vellinga (University of , Berkeley, USA), Dr. Lorenzo Pecoraro (Tsinghua University, Shenzhen, China), Dr P. Brandon Matheny, (Department of Ecology and Evolutionary Biology, University of Tennessee, Knoxville, USA), Dr. Slavomír Adamčík (Institute of Botany, Slovak Academy of Sciences), Dr. T.K. Arun Kumar (The Zamorin's Guruvayurappan College, Kerala, India), Dr. Shaun

Pennycook (Research Associate, Landcare Research Private Bag 92 170, Auckland 1142, New Zealand), Dr. Zai-Wei Ge (Kunming Institute of Botany, Chinese Academy of Sciences, P. R. China) and Dr. Kanad Das (Botanical Survey of India, Ministry of Environment, Forests and Climate Change, Cryptogamic Unit, P.O. Indian Botanic Garden, Howrah 711103, W.B., INDIA) for their valuable comments and suggestions to finalize the names of taxa and for being reviewers of the manuscripts.

I wish to thank Prof. Dr. Habib Ahmad TI (Vice Chancellor, Hazara University, Mansehra, Pakistan) and Prof. Dr. Abdur Rashid (Department of Botany, University of Peshawar, Peshawar, Pakisan) Dr. Muhammad Fiaz (Assistant Professor, Hazara University, Mansehra, Pakistan), Dr. Rosanne Healy (Postdoctoral Fellow, University of Florida, USA) for collaborative work. I must thank Genevieve E. Tocci (Curatorial Assistant, Harvard University Herbaria) and Michaela Schmull (Research and Curatorial Associate Harvard University) for helping me getting access to herbarium collections, Dr. Teresa Iturriaga for her guidance in morphological work and chemical reactions during microscopy, Mr. Danny Haelewaters, and Mr. Jason for arrangements of field tours. It gives me pleasure in writing the name of Dr. Xu Feng and Dr. Muhammad Ilyas for always being there in troubleshooting. Thankfulness is also due to Mrs. Saima Javed (Curatorial Assistant) for being so compassionate. I am not neglecting to acknowledge Mr. Shah Hussain and Dr. Chang-lin Zhao for their help in the lab and field work. Thanks to all of them for nice company at Harvard University. Special thanks to Ms. Hira Bashir for joint efforts to get the molecular results. In addition, many thanks to Mr. Ishtiaq Ahmad for helpful discussions in identification of the taxa, and assistance in the field tours. Thanks is also due to Dr. Shujaul Mulk Khan (Assistant Professor, Quaid-i-Azam University Islamabad, Pakistan) for his help in multivariate analysis. I am thankful to all my lab fellows especially Ms. Amna Imran, Mrs. Arooj Naseer, Ms. Memoona Khan, Ms. Munazza Kiran, Mrs. Fauzia Aqdas, Ms. Ayesha Zulfiqar, Ms. Rukhma Fatima, Ms. KhushBakht, Ms. Annum Razzaq, Ms. Ayesha Farooqi, Ms. Fatima Razaq, Ms. Lubna Choudry, Mr. Muhammad Usman, Mrs. Afshan Wahab, Mr. Ziaullah and my class fellow Ms. Madiha Sadiq for their joyous company during this venture.

It will be a shear neglect on my behalf, if I fail to give credit to parascientific staff and administration of the department. I highly acknowledge my friends especially Ms. Aneela Yasmeen, Dr. Ayesha Bibi, Dr. Shaista Bashir, Dr. Nousheen Yousaf, Ms. Aamna Ishaq and Ms. Tasleem Javed for their help, affection, immense consolations and wonderful company during the completion of this venture. My heartiest thanks go to my parents and siblings for their unconditional love, continual support and encouragement not only in my studies but also in my life. I am indebted to DPCC (Doctoral Program Coordination Committee), University of the Punjab, Pakistan and HEC (Higher Education Commission), Pakistan for funding this research project and awarding me the scholarships under Indigenous PhD Fellowships for 5000 Scholars, (Phase-II) and IRSIP (International Research Support Initiative Program).

Sana Jabeen

1

INTRODUCTION

Cedrus deodara (Roxb. ex D. Don.) G. Don. commonly known as Himalayan cedar or (in Urdu) is an important member of Pinaceae, an extant family of gymnosperms represented by 11 genera and approximately 235 species (Ge et al., 2012). The tree is native to Western Himalayan region of Afghanistan, China, India, Nepal and Pakistan. It is adapted to mountainous climates and found to flourish at 1500–3200 meters above sea level (Farjon, 1990). It is among the most frequent (49.6%) in Himalayan moist temperate forests of Pakistan, followed by A. B Jackson (46%) and Royle (43%) (Champion et al., 1968; Siddiqui et al., 2013). Cedrus deodara is also designated as a symbol of national tree of Pakistan (www.pakistan.gov.pk).

In Hindukush and Himalayan forests of Pakistan, Cedrus deodara has been found as a leading dominant tree species in different localities (Ahmed et al., 2011). Pure stands of this tree have been recorded from Northern Pakistan (Ahmed et al., 2010). Basically the tree has adapted to dry temperate climate; however isolated stands also occur in moist temperate (Champion et al., 1965) and sub-alpine areas (Ahmed et al., 2006). Most of these forests fall between dry and moist temperate zones of Pakistan without any sharp distinction (Hussain & Ilahi, 1991). Since the species is considered as that of dry temperate area, its presence in moist temperate regions indicates its wide ecological amplitude (Siddiqui et al., 2013).

Himalayan cedar has been proven to have great pharmacological properties with a great utility and usage as conventional medicine (Gupta et al., 2010). It has long been used in eastern medicines and its wood extract is carminative, diaphoretic and useful in fever, flatulence, pulmonary and urinary disorders (Baquar, 1989). Several alkaloids isolated from different parts of tree have been found very useful in medicine (Zhang et al., 1990; Digrak et al., 1999; Shinde et al., 1999; Rawat et al., 2000; Wolff et al., 2001; Dimri & Sharma, 2004). It has been grown as an ornamental tree, and is in great demand as building material due to its durability and rot resistance. Cedar oil is used in soap perfumes, sprays, floor polishes insecticides and microscope work as a clearing oil (Tandan et al., 1998).

Besides great economic importance of Cedrus deodara, it is an important

2 phytobiont for many ectomycorrhizal fungi (Wang & Qiu, 2006; Vaario et al., 2006; Hibbett & Matheny, 2009). Its large trunk and solid vigor is supported by belowground ectomycorrhizal fungal communities. These ectomycorrhizal fungal communities provide a balanced reciprocal association between the roots of higher plants and fungi (Frank, 1885). This symbiotic association is beneficial in natural conditions and play a major role in the functioning, maintenance and evolution of and stability as well as productivity of the ecosystem (Melin, 1923; Egger & Hibbett, 2004; Smith & Read, 2008).

The association of macro fungi in the form of ectomycorrhizal symbiosis, is of vital importance for tree growth and survival. Moreover, the establishment of the symbiosis is required for the completion of the fungal life cycle. The success of this symbiosis is mainly based upon the mutual exchange of nutrients between the symbionts. The fungi enhance host nutrient acquisition, protect against root diseases and mitigate the effect of abiotic stresses caused by chemicals, herbivores, pathogens, or drought (Colpaert, 2008; Finlay et al., 2008; Smith & Read, 2008).

To study the ectomycorrhizal fungal communities, the first step is to characterize and identify the ectomycorrhizal fungi. These fungi can be identified by their reproductive structures but these reproductive structures are not always representative of all the taxa on root tips in the form of ectomycorrhize (Visser, 1995; Gardes & Bruns, 1996; Durall et al., 1999; Jonsson et al., 1999). For the clear picture of ectomycorrhizal fungal communities, it is required to study these fungi from reproductive structures as well as ectomycorrhizal tissue.

Over the last 300 years, morphological characters, which are relatively easy to observe and record, have been used to classify fungi (Talbot, 1971). However, these characters are not supportive to identify all the fungal species, especially the mycorrhizae. Fungi that cannot be distinguished morphologically because of their intergradal color variations and intraspecific anatomical variations due to environmental factors can be separated using molecular based techniques (Eberhardt et al., 1999). In addition, morphological characters may not reflect the phylogenetic relationships due to plasticity, parallelism, and reversal (Judd et al., 2002).

During recent decades, the use of molecular data, especially DNA sequences have

3 revolutionized the process of classification allowing the reconstruction of the phylogenetic relationships leading to a renaissance in taxonomic study (Seifert et al., 2011). Morphological studies formed the base for , but recent classification systems increasingly use phylogenetic analyses based on DNA sequence data revealed major evolutionary trends among fungi (O‟Donnell 1979; Hibbett et al., 1997; Jensen et al., 1998; Sugiyama, 1998; Moncalvo et al., 2000, 2002; Tanabe et al., 2000, 2004, 2005; Gottlieb & Lichtwardt, 2001; White et al., 2001; White, 2002, 2006; Cafaro, 2005; Seif et al., 2005; Dentinger et al., 2016). Combining morphological characters and molecular data is an efficient approach to resolve taxonomic problems.

The molecular techniques are based primarily on PCR amplification of different regions or genes as molecular markers. The ideal molecular identification employs data from multiple genes, as no single locus is universally reliable for species identification (Taylor et al., 2000) but Internal Transcribed Spacer (ITS) region of nuclear rDNA exhibits a high level of variability among fungal species and minimal variation within species (Egger, 1995; Gardes & Bruns, 1996; Horton & Bruns, 2001; Kårén et al., 2008; Tedersoo et al., 2011; Hanif et al., 2012; Razaq, 2013; Yousaf et al., 2013). However, there are groups in which ITS does not discriminate between closely related species, so the use of only ITS for species recognition is not recommended (Peay et al., 2008).

In ecological studies, multi locus knowledge of species is rarely available; so multi gene approach becomes problematic because the connections between loci are lost when the sequences are generated from soil or plant material. For these reasons, ITS region of the nuclear ribosomal RNA gene, has become widely used for species identification (Horton & Bruns, 2001; Porras-Alfaro et al., 2011; Schmidt et al., 2013; Zhang et al., 2016; Long et al., 2016).

In comparison with other molecular markers used for the identification, ITS is accepted as universal DNA barcode marker for fungi (Schoch et al., 2012). It has also been used for plant phylogenetics (Buchheim et al., 2011). The value of species level match is mostly considered 97% or 98% between an unknown sample sequence and sequences from GenBank database (Tedersoo et al., 2003; Cline et al., 2005; Horton et al., 2005; Izzo et al., 2005; Morris et al., 2008; Ilyas, 2013; Long et al., 2016).

In this research work, different cedar dominating coniferous forests of Pakistan

4 were selected for the collection of ectomycorrhizal fungi. The objective of this research work is to study the ectomycorrhizal fungal communities by the characterization of taxa from the above ground in the form of fruit bodies and below ground in the form of ectomycorrhizal association. The identification is based on morphological characters combining with the use of different molecular markers to ensure the authenticity of the species identification.

This contribution focused on the parameters for community studies in terms of abundance, diversity and dominance with respect to the climatic and edaphic factors to describe the distribution patterns of ectomycorrhizal fungi. Investigations of ectomycorrhizae from the selected sites would highly support the ecological studies in future. Commercial use of ectomycorrhizal fungi could greatly improve the forest ecosystem. This research work will serve as baseline data for research in cedar forest management projects in Pakistan.

5

LITERATURE REVIEW

The ectomycorrhizal symbiosis represents one of the most prominent and ecologically crucial mutualistic associations in terrestrial habitats (Rinaldi et al., 2008). The association establishes with the fine roots of autotrophic trees and shrubs, predominantly of the families Betulaceae, Dipterocarpaceae, Fagaceae, Juglandaceae Pinaceae and Salicaceae (Watling & Lee, 1995; Béreau et al., 1997; Moyersoen et al., 2001; Henkel et al., 2002; Haug et al., 2004; Nataranjan et al., 2005; Peay et al., 2010).

Below ground symbiotic relationships between fungi and plants are continuously being discovered. It has been estimated that around 25000 fungal species are ectomycorrhizal in association with 8000 plant species (Rinaldi et al., 2008), though the number is likely to be much higher (Tedersoo et al., 2010). More ectomycorrhizal fungal species are being added to the fungal flora of the world with the increase in studies on ectomycorrhizae (Avis et al., 2008; Niazi et al., 2009; Nieto & Carbone, 2009; Niazi et al., 2010; Smith et al., 2011; Bahram et al., 2011, 2012; Ashraf et al., 2012; Hanif et al., 2012; Jabeen et al., 2012, 2014, 2015a & b, 2016a; Sarwar et al., 2013; Jabeen & Khalid, 2014; Smith et al., 2013; Seress et al., 2015).

A comprehensive account of ectomycorrhizal plant species is given by Brundrett (2009). According to him, 145 genera belonging to 26 families of plants are ectomycorrhizal. Among these 285 species belong to gymnosperms, predominates are boreal coniferous trees belonging to Pinaceae. The diversity of the fungi in an area does not follow patterns of plant diversity. For instance, coniferous forests growing in northern latitudes have hundreds of ectomycorrhizal fungal species where only a few ectomycorrhizal plant species dominate (Allen et al., 1995).

Characterization and identification of the ectomycorrhizal fungi is one of the most fundamental requirements for an understanding of ectomycorrhizal functioning in natural forest ecosystems (Brand, 1992). Recognition of huge diversity of these fungi began in the 19th century. Structural diversity of ectomycorrhizal fungi was first time revealed from the detailed drawings by Gibelli (1883) followed by Frank (1885). The first detailed descriptions at species level on morphological and anatomical basis originated from 1960s (Schramm, 1966; Chilvers, 1968). Since then many short and several detailed

6 descriptions have been published (de-Roman et al., 2005).

The study of ectomycorrhizal associations has been based mainly on the above ground reproductive structures (Molina et al., 1992; Trappe, 1962). Although there were a few pioneer studies on the classification of ectomycorrhizal roots in which only the morphological characters were used for identification (Chilvers, 1968). Dominik (1969) tried to establish classification systems in order to develop identification keys. Zak (1973) stated that a detailed description of each ectomycorrhiza was essential for identification and that this description should be based on several collections in order to note possible variations over different developmental stages.

Agerer (1987–1995) produced first comprehensive data on the basis of morphotyping. In the last few years more papers have been published concluding that morpho-anatomical features are helpful for delimitation and recognition of fungi at different systematic levels (Agerer et al., 1996, 1998; Agerer & Beenken, 1998; Agerer & Rambold, 1998; Agerer, 1999a; Eberhardt et al., 2000; Eberhardt, 2002; Agerer & Iosifidou, 2004; Beenken, 2004a & b).

Classically, the identification of ectomycorrhizal fungi in soil has been carried out by tracing the rhizomorphs (Masui, 1926). More recently, morphotyping can result in a reasonable representation of mycorrhizal fungal diversity as it relies on assessing belowground fungal structures often missed in surveys. Morphotyping allows these fungi to be traced to specific host plants and can increase the accuracy in determining associations between these fungi and their hosts (Dahlberg, 2001).

Agerer (1986, 1987–2002, 1994, 1999b) published guidelines for description and identification of ectomycorrhizae widely. In 1998, Agerer & Rambold developed a user friendly computer based System for characterization and determination of ectomycorrhizae. In addition, Agerer (1987–2002) created a system for those ectomycorrhizae described but not yet identified, edited a Colour Atlas of Ectomycorrhizae with photographs to facilitate identification by comparison.

Ingleby et al. (1990) developed a system to name the features of ectomycorrhizae, and published 24 brief descriptions including photographs and drawings. Goodman et al. (1996–2000) developed methodology, so-called Concise Descriptions of North American , giving more detailed descriptions than those by Ingleby et al. (1990),

7 but they lack the level of detail of the system proposed by Agerer (1986, 1987–2002, 1994, 1999b) and are confined to the North American geographical region.

Based on morphotype and surveys, ectomycorrhizal fungal species known to coexist in species rich communities. Numerous studies have drawn conclusions about the diversity and abundance of these fungi only from collected sporocarps (Dahlberg, 2001). However, a high frequency of one fungal species above ground does not imply below ground dominance due to the fact that the amount of fungal biomass allocated to reproductive structures varies among fungal species. Often the diversity of ectomycorrhizal fungi in an area is poorly represented above ground due to different fruiting strategies for individual species (Gardes & Bruns, 1996).

In order to characterize accurately the ectomycorrhizal fungal community in a given forest, both above and below ground fungal components must be sampled due to inconsistent fruiting strategies for some species and reliance on abiotic factors for fruiting (Gardes & Bruns, 1996). It is ideal to sample both above and below ground components of ectomycorrhizal fungi at different sites to accurately characterize the fungal taxa and to describe the community structure.

It has been observed that most of the fungal taxa either in the form of fruitbodies or mycorrhizae, closely related species often have similar morphological characters (Comini & Pacioni, 1997; Zambonelli et al., 1997, 1999; Arora & Nguyen, 2014). Over the last 25 years, the use of molecular markers to identify these fungi has greatly increased the number of known taxa (Weiss et al., 2004; Latha & Manimohan, 2016; Seress et al., 2015; Jabeen et al., 2016b). The advent of these molecular tools has opened new pathways to the study of fungal communities (White et al., 1990; Bruns et al., 1991; Gehring et al., 1998; Ilyas, 2013; Long et al., 2016). In addition, many fungal groups previously considered to be saprobic have been found to be ectomycorrhizal (Kõljalg et al., 2000). These DNA based molecular techniques are efficient tools in fungal systematics, but the morphological and anatomical features of ectomycorrhizae are still an important source of information for the better understanding of the fungal component (de- Roman et al., 2005).

Polymerase chain reaction (PCR) amplification of the Internal Transcribed Spacer (ITS) region of neclear ribosomal DNA (nrDNA) have been used to identify

8 ectomycorrhizal fungi (Gardes & Bruns, 1993; Egger, 1995). This specific region varies among species and highly conserved within species (Bruns et al., 1991; Gardes & Bruns, 1993; Egger, 1995). Thus, this approach is well suited for the identification of fungi from ectomycorrhizal roots at the species level. The ectomycorrhizal status of many additional fungal genera was discovered by comparing fungal DNA from root tips with those of fruit bodies (Gehring et al., 1998; Palfner & Agerer, 1998a & b; Tedersoo et al., 2006; Smith et al., 2007). If the fruit body and root tip sequences displayed high sequence similarity, these taxa are considered molecularly supported as ectomycorrhizal fungi. Extensive descriptions and analyses of mycorrhizae now enable about 100 species, generally from one host, to be identified (Gronbach, 1988; Ingleby et al., 1990). However, a high level of skill is required by the analyst and many morphotypes in field studies still remain unidentified (Egli et al., 1993; Kårén & Nylund, 1996; Yamada & Katsuya, 1996).

Internal transcribed spacer has been accepted as a universal barcode marker for fungi. Among the regions of the ribosomal cistron, the ITS region has the highest probability of successful identification for the broadest range of fungi, with the most clearly defined barcode gap between inter and intraspecific variation. It is easy to amplify even from small quantities of DNA due to the high copy number in the genome. About 172,000 full length ITS sequences from fungi are available in GenBank, representing almost 15,500 species and 2,500 genera (Schoch et al., 2012). Currently (May 2016), 24835 sequences from Ascomycetes, 21901 from Basidiomycetes, 844 from Zygomycetes and 3412 available in UNITE database (https://unite.ut.ee/).

Besides ITS, 18S nuclear ribosomal small subunit (SSU) and 28S nuclear ribosomal large subunit rRNA gene (LSU) combining with ITS have also been used to discriminates species (Taylor et al., 2008; Geml et al., 2009). Protein coding genes are also widely used in fungal systematics for phylogenetic analyses. Available primers for such markers usually amplify a narrow taxonomic range e.g., translation elongation factor 1-α and β-tubulin are generally used for mold genera (Schoch et al., 2009). Among protein coding genes, the largest subunit of RNA polymerase II (RPB1) could be a potential fungal barcode but it is ubiquitous and single copy in the genome, and has a slow rate of sequence divergence (Tanabe et al., 2002).

9

The work in Pakistan regarding macro fungi and their ectomycorrhizae from Pakistan has been started in the recent years. Hundreds of ectomycorrhizal morphotypes have been published in identified and unidentified form from Pakistan (Khalid & Niazi, 2003; Afshan et al., 2004; Kazmi et al., 2004; Niazi et al., 2006, 2007, 2008, 2009, 2010; Niazi, 2008; Sarwar et al., 2011, 2012, 2013; Ashraf et al., 2012; Hanif, 2012; Hanif et al., 2012; Jabeen et al., 2012, 2014, 2015a & b, 2016a; Ilyas, 2013; Jabeen & Khalid, 2014). Detailed work on ectomycorrhizal fungal communities has been conducted by Ilyas (2013) associated with deciduous trees.

This research project deals with the study of ectomycorrhizal fungi associated with Cedrus deodara (Roxb. ex D. Don) G. Don. The occurrence of natural ectomycorrhizal association with this plant species is virtually unexplored in the form of detailed community study. Only a few reports of ectomycorrhizal association with C. deodara have been published from (de-Roman et al., 2005) and Asia (Sharma & Singh, 1990; Nizzar-Hocine et al., 1998; Singh & Lakhanpal, 2000; Bisht et al., 2003; Hall, et al., 2003; Niazi et al., 2006; Niazi, 2008; Deepika et al., 2011; Hanif et al., 2012; Kumari et al., 2012; Sarwar et al., 2012). The important mycorrhizal symbionts of C. deodara are several species of Amanita Pers., Boletus Tourn., Craterellus Pers., Pers. and Russula Pers. (Sharma & Singh, 1990; Nezzar-Hocine et al., 1998; Singh & Lakhanpal, 2000; Bisht et al., 2003; Hall et al., 2003; Kumari et al., 2012).

Species abundance, diversity, and dominance are important parameters to determine the community structure of ectomycorrhizal fungi. These parameters depend upon the climatic, edaphic, physiognomic and physiographic conditions of the area. Mean annual temperature and precipitation, size of host plant and their distribution on the landscape also influence the community structure (Morris et al., 2008; Tedersoo et al., 2010; Bahram et al., 2011).

Present research work provides the detailed community structure of ectomycorrhizae associated with Himalayan cedar from different areas of Pakistan. The data is analyzed from both, above and below ground ectomycorrhizal community components. The study provides an understanding about the factors involved in community composition. Community structure assessment forms a sound basis for application to forestry practices and potentially inoculation programs. Correlation between sporocarp diversity and below ground ectomycorrhizae as well as between the

10 climatic and edaphic factors also provide more thorough understanding of community structure.

11

AIMS AND OBJECTIVES

The main objectives of this study include:

 To isolate and characterize the ectomycorrhizal fungi associated with Cedrus deodara.  To identify the ectomycorrhizal fungal species, those have not been recorded from above ground collections.

 To determine the overall and stand specific diversity of sporocarps and belowground communities.

 To generate reference data from local collections for future taxonomic studies.

The results obtained from the knowledge of above and below ground community structure will be helpful to decide about those taxa which can be used for future forest establishment programs.

12

MATERIALS AND METHODS

3.1. Sampling site description Pakistan has a great variety of landscapes with diversified topography. Geographically Pakistan is located within the latitude and longitude of 30° 00 N, 70º 00 E including 803, 940 km2 of total land area. It has some of the world‟s highest cold areas that occur above 5,175 meters above sea level in the Himalayas and the hottest low areas in the Indus Plains with many intermediate ecological zones (UNEP, 1998). Northern area of Pakistan covers about 72,500 km2 neighboring China is linked via the famous Silk Road through the Khunjrab Pass, adjoins the distributed territory of Kashmir to the East and Afghanistan to the West (Ahmed et al., 2011).

Himalayan, Hindu Kush and Karakoram ranges are among the most prominent features on the face of earth. Almost 60% of the natural forests resources are concentrated in these mountainous regions. Coniferous forests occur from 1,000 to 4,000 m altitudes. Abbotabad, Balochistan, Azad Kashmir, Chitral, Dir, Swat, Malakand, Mansehra and Murree are rich in Abies pindrow, Cedrus deodara, Juniperus macropoda Boiss, Pinus gerardiana Wall. ex D. Don, Picea smithiana Boiss, Pinus wallichiana and Taxus fuana Nan Li and R. R. Mill (Siddiqui et al., 2013). Cedrus deodara is commonly distributed in various locations of Abbottabad, Azad Kashmir, Chillas, Chitral, Dir, Hazara, Kaghan, Murree and Swat valley (Ahmed et al., 2010, 2011; Siddiqui et al., 2013).

Sampling was carried out from different coniferous forests coming under three administrative divisions (Rawalpindi , Hazara & Malakand) of Punjab and Khyber Pakhtunkhwa province of Pakistan (Ahmed et al., 2011) (Figure 1). Kalam and Mashkun are situated in Malakand division of Khyber Pakhtunkhwa province. These areas are occupied by offshoots of Hindu Kush range (Hamayun, 2003). These ranges are dominated by Cedrus deodara forests along with Pinus species and Quercus oblongata D. Don [= Q. incana Roxb., nom. illeg.] (Champion et al., 1965). These areas have a typical dry temperate climate (Stucki & Khan, 1999). Khanian and Khanspur come under Hazara division in KPK province, which lies immediately South of the main Himalayan range with typical moist temperate climate and is dominated by C. deodara along with Abies pindrow Royle and Pinus wallichiana A.

13

Figure 1. Map of Pakistan showing sampling sites. A. Hazara division; B. Malakand division; C. Rawalpindi division. Tree symbol showing sampling stands.

14

B. Jacks. (Siddiqui et al., 2013). Patriata and Kuzah Gali come under Rawalpindi division in Punjab province with sub-tropical and moist temperate forests dominated by C. deodara and P. wallichiana.

3.2. Sampling strategy

From Hazara division; Khanian (stand 1), and Khanspur (stand 2), from Malakand division; Kalam (stand 3) and Mashkun (stand 4), from Rawalpindi division; Kuzah Gali (stand 5) and Patriata (stand 6) were selected as sampling stands (Figure 1).

Following points were taken under consideration while selecting the stands: 1. domination of Cedrus deodara, 2. visual homogeneity of vegetation, 3. free from recent anthropogenic disturbance, 4. at least one square kilometres or more area.

3.3. Sampling of sporocarps

Sporocarps growing in the vicinity of Cedrus deodara were collected from the sampling sites. Sporocarps collected from under the one tree was considered as one sample.

3.4. Sampling and preservation of fruit bodies

Sporocarps growing in the cedar dominating forests were collected. Each collection was assigned a collection number, site and date of collection and collector‟s name. With the help of digger, atleast two individual specimens were collected; one for experimental purpose and other for voucher specimen. Other individuals were recorded for statistical purpose. Characteristics of the habitat and associated vegetation were also recorded. Adhering soil particles were cleaned off and photographed using a Nikon D70S digital camera. Macro-morphological features of the specimen were examined in the field. Colors were designated using Munsell (1975) color charts. Small tissue from the was stored in 2% CTAB buffer and the remaining was subjected to air drying for long term preservation. Large fruiting bodies were cut into longitudinal sections for effective drying.

3.5. Morphological studies of fruit bodies

For detailed morphological examination, tissues from lamellae, pileipellis and stipitipellis were mounted on glass slides and observed in different media. Phloxine (1%)

15 was used for better contrast, Melzer‟s reagent for amyloid ornamentation, and KOH (5%) for colored hyphae. Anatomical features were observed under Meiji Techno MX4300H compound microscope. Measurements were recorded using a Carl Zeiss Jena ocular micrometer and line drawing were made using a Leitz Wetzlar camera lucida.

For scanning electron microscopy, tissue from the was mounted on metal stub using double sided adhesive tape covered with a fine layer of gold by sputtering in a vacuum sputterer (JFC-1100, JEOL, Japan).

The abbreviation „n/b/p‟ was used to represent „n‟ basidiospores measured from „b‟ basidiomata from „p‟ collections. Basidiospore dimensions were given as length × width (l × w), with extreme values given in parenthesis. Q = l/w ratio of individual ; avQ = average Q of all spores ± standard deviation (Ge et al., 2010). Voucher specimens were deposited in the Herbarium, Department of Botany, University of the Punjab, Quaid-e-Azam Campus, Lahore, Pakistan (LAH).

3.6. Sampling of morphotypes

A mature tree with a trunk diameter of 50–60 cm was selected as a host for sampling of ectomycorrhizal morphotypes. Soil blocks of 15 cm3 including roots of the tree were excavated at 15 cm to 1m away from the trunk of each tree. Two blocks per each individual tree were taken at each sampling time. The blocks were placed in plastic bags, labelled with collection number, stand code, locality and date. The samples were brought to the laboratory and stored at 4oC until washing and morphotyping.

3.7. Soil sampling and analysis

An extra amount (100 g) of soil was taken from the same position where the morphotypes were sampled. All soil samples from the same stand were mixed to form a composite sample for each year. Soil analysis was performed at Soil and Water Testing Laboratory, Agriculture Department, Government of the Punjab, Lahore.

3.8. Cleaning, morphotyping and storage of ectomycorrhizae

The soil blocks were soaked in water for overnight to loosen the adhering soil particles then shifted to 2 mm sieve and placed under running water to remove the soil. The ectomycorrhizal root tips were carefully sorted into morphotypes under incandescent

16 light according to their morphological features including ramification, colour, size and emanating elements. The attached soil particles were removed with the help of fine brush under Meiji Techno EMZ-5TR stereomicroscope. Individual root tips were dissected and morphologically identical morphotypes from one sample were joined for a morphotypes sample. These morphotypes were kept in McCartney bottles in distilled water for later morphological examination. Replicates of these morphotypes were kept at 8°C in eppendorf tubes containing 1 mL of 2% CTAB buffer for molecular analysis.

3.9. Morphological studies of ectomycorrhizae

Morphological characters were noted under the stereomicroscope. For anatomical details, outer and inner mantle layers and radiating hyphae were mounted in trypan blue stain (Agerer, 1991). Hyphae were measured using an ocular micrometer and drawn using a camera lucida. Morphotypes were anatomically characterized following Agerer‟s (1999b) methodology.

3.10. Molecular studies

3.10.1. DNA extraction

Genomic DNA was extracted from fruit bodies using CTAB extraction buffer following Bruns (1995) with slight modifications. Where the CTAB method failed in extraction, DNeasy Plant Mini Kit (Qiagen) was used following manufacturer's protocol. For DNA Extraction from root tissue, firstly CTAB method was used, in case of failures, then Extract-N-Amp™ kit was used for extraction following manufacturer's protocol. In some cases, DNeasy Plant Mini Kit (Qiagen) was also used for DNA extraction from genomic DNA from ectomycorrhizal fungi and host. For visualization of DNA, gel electrophoresis was performed.

Agarose (1%) was prepared by melting of molecular grade agarose in 1x TAE buffer in microwave oven. It will be allowed to cool for a couple of minutes then 2.5 μL of ethidium bromide or gel red was added. Gel was casted and allowed to polymerize for a minimum of 20 minutes at room temperature. Each DNA sample was loaded in separate wells stained with loading dye. DNA ladder mixture was also loaded in a well for comparison. Electrophoresis was performed for 20 minutes at 90 V power and the bands were visualized under UV light in gel documentation system and photographed digitally.

3.10.2. Polymerase chain reaction (PCR)

17

Different sets of primers were used for the amplification of specific regions. For most of the taxa, especially fungi from ectomycorrhizal tissue, ITS primers ITS1-F (5'- CTTGGTCATTTAGAGGAAGTAA-3') or ITS5 (5'-GGAAGTAAAAGTCG TAACAAGG-3') along with ITS4 (5'- TCCTCCGCTTATTGATATGC-3') were used for amplification (White et al., 1990; Gardes & Bruns, 1993).

For LSU amplification LR0R (5'-ACCCGCTGAACTTAAGC-3') and LR5 (5'- TCCTGAGGGAAACTTCG-3') were used. For some species identification, rpb2 and tef- 1α gene regions were amplified using the primer combinations gRPB2-6F (5'- TGGGGKWTGGTYTGYCCTGC-3') with gRPB2-7R (5'-CCCATWGCYTGCTT MCCCAT-3') and EF1-983 (5'-GCYCCYGGHCAYCGTGAYTTYAT-3') with REF1- 1567F (5'-ACHGTRCCRATACCACCRATCTT-3'), respectively (Ge et al., 2014, 2015). PCR reaction was performed using Extract-N-Amp™ PCR kit following manufacturer‟s instructions and the products were sent to Macrogen Inc. (South Korea) for purification and sequencing. For host identification in ectomycorrhizal association, 28C (5'- GCTATCCTGAGGGAAACTTC-3') and 28KJ (5'-GGCGGTAAATTCCGTCC-3') primers were used for amplification (Cullings, 1992).

Manual PCR was performed in 25 µL reaction volume, containing 2.5 µL 10X Econo Taq Buffer, 0.5 µL dNTPs, 1.25 µL of each primer (10 µM/µL), 0.125 µL of

Econo Taq® DNA Polymerase, 14.375 µL H2O and 5 µL of DNA template. Thermal cycler conditions for amplification were the same as described by Ge et al. (2014), except that annealing temperatures were optimized for each gene region: 54ºC for ITS of fungi and pant, 52ºC for nrLSU, and 60ºC for rpb2 and tef-1α. PCR products were purified using the QIAquick PCR Purification kit (QIAGEN, Valencia, CA, USA).

In case of ectomycorrhizal tissue, sometimes 2 and 3 bands of PCR product of fungal ITS were observed on the gel. For the separation of all these products, the PCR reaction was performed in two more replicates to increase the product and then gel electrophoresis was performed using 8 % of gel with wells of 60–100 µL capacity for 2 hours at 40 V. The bands were cut under blue light transilluminator and QIAquick Gel Extraction Kit was used for separation of DNA from gel following manual.

18

Sequencing was performed with the same PCR primers using the Big Dye Sequencing Kit v.3.1 on an ABI-3730-XL DNA Analyzer (Applied Biosystems, Foster City, CA, USA). For the removal of excess DyeDeoxy terminators, centri-sep protocol was performed prior to sequencing following Princeton Separations centri-sep spin columns user's manual. Some PCR products were also sent to Macrogen, Inc. South Korea for sequencing.

3.10.3. Sequence assembly and data mining

Consensus sequences were generated from the sequences obtained by the forward and reverse primers in BioEdit sequence alignment editor version 7.2.5 (Hall 1999) and then BLAST searched at NCBI (http://www.ncbi.nlm.nih.gov/) and UNITE (https://unite.ut.ee/). Taxa collected in the form of ectomycorrhizal fungal tissue and their fruiting structures were not found from the above ground survey were considered as operational taxonomic units (OTUs) with ≥ 97% sequence similarity and without taking into account the differences in sequence length were considered the same taxon (Arnold et al., 2007). The taxa as OTUs with <97% similarity in BLAST search were designated with running code numbers for the specific epithet until their fruit body has been found in further studies.

3.10.4. Multiple alignments and phylogenetic analysis

Sequences with closest match were selected from GenBank to reconstruct phylogeny. The sequences which showed less query cover and negative E value were left aside. Published sequences of the closest relatives of the species were also included to reconstruct phylogeny. GenBank and Unite accessions were given with each taxon name in the phylogenetic tree. For rooting of tree, near relative of the taxa involved in the tree was chosen as out group. Multiple sequences were aligned using online MUSCLE tool at EMBL-EBI (http://www.ebi.ac.uk/). Maximum likelihood (ML) analysis was performed in MEGA version 6 (Tamura et al., 2013) software to test the phylogeny at 1000 bootstrap replicates. Percentage identity and divergence in nrDNA-ITS were analyzed using MegAlign (DNASTAR, Madison, WI).

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3.11. Climatic data collection

Climatic data of the sampling areas during the sampling years was collected from Pakistan meteorological department, Headquarter Office Sector H-8/2 Islamabad.

3.12. Statistical analysis

For community analysis, data from all the stands were compiled and categorized as above ground and below ground taxa. Several parameters were taken into consideration for detailed community study. The diversity over spatial scales was described in terms of alpha, beta, and gamma diversity (Whittaker, 1972). Alpha diversity refers to the diversity within a stand and expressed by the number of species. Beta diversity was measured to examine the variations in species diversity between the major regions for comparison. Gamma diversity is the geographic scale species diversity which measures the overall diversity of ectomycorrhizal fungi associated with Himalayan cedar in all the studied sites.

For alpha diversity analysis, different indices were calculated to describe the distribution of taxa among the stands and their status in terms of species abundance, dominance and richness. Simpson's Index (D) of diversity is a quantitative measure of probability that two individuals randomly selected from a sample will belong to the same species. Simpson's Dominance Index is the simpler form of D that gives information about the dominant species. The Shannon diversity index (H) is another commonly used index to characterize species diversity in a community. It accounts for both abundance and evenness of the species present in a stand (Magurran, 2013).

Margalef's index (d) is a sub estimation of these two indices (Gamito, 2010). It measures species richness and it is highly sensitive to sample size although it tries to compensate for sampling effects (Margalef, 1958; Magurran, 2013). Margalef's index was used in combination with Berger-Parker which is sensitive to evenness or changes in dominant species (Berger & Parker, 1970).

Menhinick's index is a particular measure of species richness based on the number of samples and the expected number of species (Menhinick, 1964). Buzas & Gibson‟s index (E) is another parameter to measure the evenness depending upon the number of samples taken from each stand (Hayek & Buzas, 2010). Gini's coefficient is a

20 measure of inequality. It was used to measure any form of uneven distribution of taxa in each sample within a stand (Gini, 1912). Brillouin's index (H) is the measure of equitability in each stand (Pielou, 1969, 1975; Maurer & McGill, 2011). Formulae used for the calculation of all these indices are enlisted in Table 1. To rank the species according to their abundance, rank abundance curves were plotted according to Magurran (2013).

Beta diversity was measured in terms of Jaccard's index and Bray-Curtis. Jaccard's index was used to compare the similarity and diversity of the data collected from each stand (Jaccard, 1901). Bray-Curtis was used to quantify the compositional dissimilarity between two different sites, based on counts at each stand (Bray & Curtis, 1957). One way ANOVA was used for the analysis of variance between the stands (Fisher, 1918). Canoco software package (version 5) was used for multivariate data analysis (Šmilauer & Lepš, 2014). Using EstimateS (version 7.5) program, accumulating species richness in all major regions were visualized (Colwell et al., 2004). Chao2 species richness estimates were also calculated to infer the completeness of sampling at each site. All the data from each stand at major region was combined to analyze gamma diversity of ectomycorrhizal fungi.

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Table 1. List of formulae used for the calculation of diversity indices.

Index Formula

( ) Simpson Index = ( )

Dominance Index= ( ) ( )

Berger-Parker Dominance Index=

∑ * ( )+ Shannon Index=

Margalef Richness Index=

Menhinick Index= √

* ( )+ Buzas & Gibson's Index=

* ( )+ Brillouin's index =

22

RESULTS

During the field survey, 262 ectomycorrhizal fruit bodies were observed and identified. Total 919, morphotypes were isolated from the soil. Atleast one individual from the fruit bodies of each collection and some morphotypes from each sample were passed through molecular process. Fruit bodies found to be ectomycorrhizal in BLAST seach were then chacterized morphologically and anatomicallyalond with molecular phylogenetic analysis. Those taxa which were proven to be non ectomycorrhizal by consulting the literature are enlisted in Table 2. Some seprobic taxa found from the fruitbodies during molecular analysis are given in Table 3. Below ground taxa found to be identical to the fruit bodies on molecular basis, were described morphologically and anatomically. Others are enlisted in Table 4 as OTUs. During the molecular analysis of morphotypes, some endosymbionts and seprobic taxa and those forming mycorrhizal associations other than ectomycorrhizal association were identified. These taxa are enlisted in Table 5. From each stand, some ectomtcorrhizal morphotypes were selected for host identification based on molecular phylogenetic analysis. The fungi found in association with host species other than Cedrus deodara were left aside from the final data. The list of host taxa identified during this investigation is given in Table 6. Stand wise results of different soil analysis parameters are indicated in Table 7. Published data is attached in annexure.

4.1. CHARACTERIZATION OF ABOVE GROUND ECTOMYCORRHIZAL FUNGI

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Genus Amanita

Amanita ahmadii nom. prov. (Figures 2 & 3)

Etymology: The specific epithet refers to the name of pioneer mycologist Sultan Ahmad (Late) from Pakistan.

Pileus upto 4 cm broad, convex to flat at maturity; cuticle off white (2.5PB9/2) when young becoming gray (2.5BG4/2) with time; areolate with abundant squamules. Squamules gray (2.5Y4/2) to dark blue (7.5PB1/2); surface dry; margins smooth, incurved and rimose at maturity. Lamellae off white (2.5BG4/2) to cream (5Y9/4) up to 0.3 cm broad, adnexed, sub distant to close, edges entire. Lamellulae small (2/6 of the lamellae), single tiered. 6.7 × 0.6 cm, apex slightly wide up to 0.9 cm in diameter, bulbous base up to 1.5 cm in diameter, central, cylindrical, surface smooth to fibrillose, white (2.5BG4/2) to cream (5Y9/4) towards the apex and grayish brown (5GY5/2) towards the base.

Basidiospores [50/2/1] (4.2) 4.9–8 (8.5) × (4.2) 4.3–7.3 µm, Q = (1.33) 1.22–1.03 (1), avQ = 1.10 ± 1.96; globose to subglobose, exine smooth, apiculus prominent, grayish in 5% KOH. Basidia (67.3) 59–34.7 (32.3) × (8) 7.8–7 (6.8) µm, clavate having 2–4 starigmata, thin walled, wall up to 3 µm thick, hyaline 5% KOH. Cystidia (30.9) 28.5– 25.2 (24.8) × (7.5) 7.3–7 (6.8) µm, clavate, hyaline in 5% KOH. Partial veil remnants hyphae bear terminal sub globose to elongated cells (56.6) 54–49.5 (42.5) × (19.1) 16– 13.4 (13) µm on a branched filament (4) 3.8–3.5 (3.3) µm wide, septa frequent, clamp connections not observed. Pileipellis hyphae globose, sub globose to elongated (43.2) 33– 25.6 (24) × (24.3) 20–16.5 (16) µm with a filament (4.7) 4.2–4 (3.7) µm in diameter, branched, septate, clamped septa frequent, light brown with some hyaline tissue in 5% KOH. Squamules hyphae globose to sub globose (12.7) 12.2–8 (6.8) × (10.8) 10.6–7.5 (4.4) µm with a filament (3.5) 2.6–0.9 (0.7) µm in diameter, mixture of brown and hyaline cells in 5% KOH. Stipitipellis hyphae (24) 22.6–3.5 (3) µm wide, filamentous, branched, broad terminals, hyaline in 5% KOH, septate, clamp connections not observed.

Material examined: Pakistan, Khyber Pakhtunkhwa province, Malakand division, Swat district, Mashkun, 2500 m asl, on soil under Cedrus deodara (Roxb. ex D. Don) G. Don, 5 Sep 2013, Sana Jabeen K4-38; SJ35 (LAH35010).

KR674106_Cortinarius_albidolilac KR674105_Cortinarius_albidolilac 99 KR674104_Cortinarius_albidolilac KR674102_Cortinarius_albidolilac 71 KR674103_Cortinarius_albidolilac EU057055_Cortinarius_lilacinovel EU655668_Cortinarius_lilacinovel 77 99 EU056960_Cortinarius_lilacinovel 70 95 EU056959_Cortinarius_lilacinovel 99 KF673472_Cortinarius_cobaltinus KF673471_Cortinarius_cobaltinus 17 96 EU056969_Cortinarius_spectabilis 93 DQ663241_Cortinarius_caesiocinct 98 EU056978_Cortinarius_calochrous 95 DQ663234_Cortinarius_calochrous EU056977_Cortinarius_calochrous KF732347_Cortinarius_metarius 89 42 EU057053_Cortinarius_barbarorum EU655671_Cortinarius_barbarorum 99 DQ663236_Cortinarius_barbarorum DQ663238_Cortinarius_barbarorum Calochori EU057059_Cortinarius_largentii DQ323964_Cortinarius_corrosus EU056974_Cortinarius_corrosus 80 Cortinarius_corrosus SJ41 FJ039642_Cortinarius_corrosus KF732272_Cortinarius_calojanthin 76 FJ039641_Cortinarius_corrosus DQ663281_Cortinarius_corrosus KF732538_Cortinarius_calojanthin EU056975_Cortinarius_corrosus EU057057_Cortinarius_corrosus KF732561_Cortinarius_sublilacino 99 EU655674_Cortinarius_sublilacino EU056988_Cortinarius_sublilacino EU056987_Cortinarius_albovestitu 59 62 EU057056_Cortinarius_haasii EU655684_Cortinarius_aurantioruf 99 AY669561_Cortinarius_haasii EU056985_Cortinarius_haasii 67 EU056986_Cortinarius_haasii AY174787_Cortinarius_glaucopus Out group

0.01

24

Molecular phylogenetic characterization (Figures 4 & 5)

Sequencing of the PCR products of ITS region of Amanita ahmadii yielded 712– 714 base pairs using ITS1F and ITS4 primers. Consensus sequence of 615 base pairs was obtained by trimming the motif. When BLAST searched at NCBI, A. ahmadii showed 98% identity to sequences from China (KJ466372 & KJ466373) with 94–100% query cover. It also showed 95% identity with sequence from Bulgaria (JX515561) with 100% query cover and 0.0 E value. It also showed 99% identity to JF908673 with 93% query cover and 0.0 E value.

Sequencing of the PCR products of LSU yielded 1175–1220 base pairs by using LR0R and LR5 primers. Consensus sequence of 876 base pairs was BLAST searched at NCBI. The subject sequence showed 99% identity to sequences from South Korea and USA (KF245897, KT072738 & HQ539663) with 99% query cover and 0.0 E value.

To reconstruct phylogeny, closely related ITS and LSU sequences were retrieved from the GenBank. Taxa from subgenus Lepidella (E.-J. Gilbert) Veselý, section Phalloideae (Fr.) Quél. were chosen as out group. The sequences generated during this study clustered with the similar taxa in section Validae (Fr.) Quél, of subgenus Lepidella. Amanita ahmadii clustered with A. aff. fritillaria ZLY-2014, A. franchetii (Boud.) Fayod and A. aspera sensu auct. mult. forming a sister with A. spissa (Fr.) P. Kumm. and allies with 66% boot strap value in ITS dataset. The species was separated from A. aff. fritillaria with a strong bootstrap value of 94% and 68% in ITS and LSU sequence analyses, respectively.

Comments

Amanita ahmadii nom. prov. is characterized by its white to gray pileus surface with abundant bluish squamules and having rimose margins. Anatomically it is characterized by its globose to subglobose basidiospores. Amanita fritillaria Sacc. morphologically differs from A. ahmadii by its grayish brown pileus and ellipsoid spores (Sanmee et al., 2008; Kim et al., 2013). Amanita franchetii and A. aspera, recently renamed as A. augusta Bojantchev & R.M. Davis (Borjantchev & Davis, 2012) is also distinct morphologically from A. ahmadii by its ellipsoid spores and yellowish brown pileus with yellow squamules. Amanita aff. fritillaria, a closely related taxon forms a sister clade with A. ahmadii. Amanita ahmadii is distinct on molecular basis as well as

25 morphological characters in having brownish and purplish in comparison with collections by Cai et al. (2014).

26

Figure 2. Morphology of Amanita ahmadii. A–D. Basidiomata LAH35010 (holotype). Bars: A = 0.8 cm; B = 1.5 cm; C & D = 1 cm.

27

Figure 3. Anatomy of Amanita ahmadii. A–F. LAH35010. A. Basidiospores; B. Basidia and Cystidia; C. Pileipellis; D. Squamules; E. Partial veil; F. Stipitipellis. Bars: A = 0.8 µm; B = 90 µm; C = 40 µm; D = 18 µm; E = 10 µm; F = 22 µm.

28

KJ638283_A_flavoconia KJ638282_A_flavoconia KM373250_A_flavoconia JF313657_A_flavoconia 98 GQ415317_A_flavoconia JN020966_A_flavoconia JF313655_A_flavoconia KJ638281_A_flavoconia 100 GQ415317_A_flavoconia JN020967_A_flavoconia JF313655_A_flavoconia KF937301_A_flavoconia 52 FJ890029_A_flavoconia KF937300_A_flavoconia 100 KF937299_A_flavoconia KP284300_A_morrisii 20 KP284299_A_morrisii KR919760_A_morrisii 100 KR919762_A_morrisii KT213441_A_morrisii 26 KF245911_A_flavipes KC185406_A_flavipes 100 AY436455_A_flavipes FJ375327_A_flavipes 70 AF085486_A_spissa AM084702_A_rubescens 100 AY436453_A_excelsa 30 60 JF907760_A_rubescens 64 AJ889924_A_spissa 86 AJ889924_A_spissa 94 EF493271_A_spissa 74 EF493270_A_spissa KF245910_A_spissa KJ638284_A_rubescens 92 A ahmadii SJ35 94 A ahmadii SJ35b 66 KJ466372_A_aff_fritillaria KJ466373_A_aff_fritillaria 100 sect. Validae 34 AB0156951_A_sp 60 AF085485_A_aspera 56 JX515561_A_franchetii 100 JX515563_A_franchetii 64 JX515562_A_franchetii 100 KJ535437_A_novinupta KF561974_A_novinupta 100 KJ535438_A_novinupta GQ166902_A_flavorubescens JF313650_A_flavorubescens 62 EU819454_A_cf_flavorubesce 86JF313652_A_rubescens AJ889923_A_rubescens AM087243_A_rubescens 66 JF313652_A_rubescens FJ890030_A_rubescens 76 FJ890031_A_rubescens 86 86 JF313651_A_rubescens AJ889922_A_rubescens 46 KM409441_A_rubescens 78 KF245918_A_rubescens KF245919_A_rubescens 58 AB015682_A_rubescens 70 100 KM052530_A_rubescens 76 KM052535_A_rubescens JF273507_A_rubescens FJ890033_A_brunneoloculari 92 EU819464_A_rubescens 60 JN020972_A_rubescens GQ250403_A_novinupta 90 KC152065_A_novinupta 90 KC152067_A_novinupta 94 KC152066_A_novinupta JF273505_A_fritillaria AY436457_A_fritillaria 100 100 JF899548_A_porphyria AB015677_A_porphyria 100 AB015679_A_citrina KC755034_A_phalloides KJ638292_A_bisporigera sect. Phalloideae 100 EF493032_A_virosa 36

0.05 Figure 4. Molecular phylogenetic analysis of Amanita ahmadii based on ITS sequences. The evolutionary history was inferred by the Maximum Likelihood method using General Time Reversible model. The analysis involved 79 nucleotide sequences. There were a total of 718 positions in the final dataset. Sequences generated from LAH35010 are marked with .

29

61 HQ539735_A_rubescens FJ890047_A_sp 46 FJ890044_A_brunneolocularis FJ890043_A_rubescens AF097383_A_rubescens AF042607_A_rubescens 58 KP711840_A_sp AF097382_A_rubescens KP711837_A_sp KP711838_A_sp_ 34 KF245903_A_rubescens 70 KF245902_A_rubescens 88 HQ539693_A_flavoconia 78 EU522816_A_flavoconia_ FJ890041_A_flavoconia 81 AF097381_A_franchetii 33 15 HQ539664_A_aff_flavorubens HQ539694_A_flavorubens 83 KF561978_A_novinupta HQ539716_A_novinupta 79 83 KJ535441_A_novinupta 24 KF245897_A_fritillaria KJ466480_A_aff_fritillaria 73 A ahmadii SJ35b 68 83 A ahmadii SJ35 AF097380_A_flavorubescens 52 AY436502_A_sp HQ539663_A_aff_flavoconia 73 AF042609_A_flavoconia KT072737_A_sp sect. Validae KT072738_A_sp KF245894_A_spissa HQ539691_A_excelsa 68 AY436491_A_excelsa 61 82 HQ539743_A_spissa_ 99 HQ539706_A_luteolovelata HQ539705_A_luteofusca KT006766_A_brunnescens KP284284_A_brunnescens 99 KP284285_A_brunnescens AF097379_A_brunnescens 66 HQ539661_A_aff_brunnescens HQ539674_A_brunnescens KP284283_A_brunnescens KR865979_A_lavendula 33 AF097378_A_citrina 88 HQ539662_A_aff_citrina 46 EU522722_A_citrina KF245892_A_citrina AF041547_A_citrina 99 AF097377_A_citrina 66 FJ890037_A_sp 99 FJ890046_A_citrina 60 HQ539679_A_citrina 74 KF245893_A_citrina AY436500_A_porphyria 96 KP866192_A_porphyria KP866189_A_porphyria 99 KP866184_A_porphyria KP866185_A_porphyria AF159086_A_virosa sect. Phalloideae

0.005

Figure 5. Molecular phylogenetic analysis of Amanita ahmadii based on LSU sequences. The evolutionary history was inferred by the Maximum Likelihood method using General Time Reversible model. The analysis involved 61 nucleotide sequences. There were a total of 874 positions in the final dataset. Sequences generated from LAH35010 are marked with .

30

Amanita brunneopantherina nom. prov. (Figures 6 & 7)

Etymology: The specific epithet refers to the brownish color of the pileus with panther like appearence due to complete veil remanents.

Pileus 3–6 cm broad, convex to plano convex or flat at maturity; cuticle golden brown (2.5Y8/4); squamules common, off white (5Y9/2); surface dry, reflective and steriate; margins entire to dentate due to striations, incurved when young becoming straght at maturity. Lamellae white to off white, up to 0.28 cm broad, adnexed, sub distant to close, edges entire. Lamellulae absent. Stipe up to 8 × 1 cm, apex slightly narrow up to 0.8 cm in diameter, base bulbous, having scales on the volva up to 2.2 cm in diameter, central, cylindrical, smooth, context off white (5Y9/2); annulus prominent membranous, skirty.

Basidiospores [40/2/2] (5.4) 6.0–9.2 (12.0) × (5.2) 6.0–8.8 (12.2) µm, smooth, globose, apiculus prominent, grayish green in 5% KOH. Basidia 50.0–67.8 × 13–15.2, clavate having 4 sterigmata, guttulate, hyaline in 5% KOH. Cystidia 28.5–30.6 × 6.5–13.3 µm, clavate, hyaline in 5% KOH. Pileipellis hyphae bears inflated globose to subglobose terminals, 47.2 x 42.5 µm , on a filament up to 11.8 µm wide hyaline in KOH. Stipitipellis 9.5 × 7.0 µm, filamentous, branched, fusiform terminals, infrequently septate, hyaline in KOH.

Material examined: Pakistan, Khyber Pakhtunkhwa province, Malakand division, Swat district, Kalam, 2400 m asl, on soil under Cedrus deodara, 4 Sep 2013, Sana Jabeen & Abdul Nasir Khalid K2-1 (LAH35011); K2-5; SJ3 (LAH35012); Mashkun 2500 m asl, on soil under Cedrus deodara, 5 Sep 2013, Sana Jabeen & Aamna Ishaq K4-29; SJ56 (LAH35014).

Molecular phylogenetic characterization (Figure 8)

Sequences of the ITS region of Amanita brunneopantherina consist of 723–726 nucleotide base pairs using ITS1F and ITS4 primers. BLAST of the consensus sequence of 623 base pairs revealed that the sequences generated during this study match 96% with (JF899547) from Canada and USA (EU525997 & EU909452) with 96% query cover and 0.0 E value.

31

For phylogenetic analysis, closely related sequences from GenBank were retrieved. Taxa from section Vaginatae (Fr.) Quél., were chosen to root the tree. The sequences generated during this study clustered with the similar taxa in section Amanita Pers. of subgenus Amanita. Amanita brunneopantherina nom. prov. was clustered in a clade having Amanita sequences from Canada and USA, but both of these taxa get separated with a strong boot strap value (63%) forming sister in section Amanita.

Comments

Amanita brunneopantherina nom. prov. is characterized by its golden brown reflective pileus and globose spores. Morphologically, the species differs from A. pantherina (DC: Fries) Krombh. by the color of the pileus and spores shape. The pileus cuticle of A. pantherina is hazel brown or dark brown while in A. brunneopantherina, it is light brown. Moreover the spores also differ in size and shape. Amanita brunneopantherina bears globose spores comparatively larger in size while A. pantherina bears small elliptical spores (Kuo, 2005). Molecular data set comparison also revealed that these two taxa are distinct from each other.

32

A

B C

Figure 6. Morphology of Amanita brunneopantherina. A–C. Basidiomata. A. LAH35012 (holotype); B & C. LAH35011. Bars: A= 2.5 cm; B = 1.5 cm; C = 6 cm.

33

Figure 7. Anatomy of Amanita brunneopantherina. A–E. LAH35012. A. Basidia; B. Basidiospores; C. Cystidia; D. Stipitipellis; E. Pileipellis. Bars: A = 14 µm; B = 5 µm; C = 7 µm; D & E = 9.5 µm.

34

56 KF651005_A_parvipantherina 58 KF651007_A_parvipantherina KF651006_A_parvipantherina 85 KF651008_A_parvipantherina AY436466_A_cf._pantherina KF650998_A_parvipantherina 62 KF651009_A_parvipantherina KM052551_A_pantherina KF017943_A_pantherina KF017944_A_pantherina 9768 KJ609156_A_pantherina KF017942_A_pantherina KF017947_A_subglobosa 66 JN943177_A_subglobosa JN943176_A_subglobosa 91 Amanita brunneopantherina SJ56 Amanita brunneopantherina SJ56b JF899547_A_pantherina 59 63 EU525997_A_pantherina sect. Amanita 100 EU909452_A_pantherina KR919757_A_breckonii 91 KJ535439_A_breckonii KR919758_A_breckonii HQ604823_A_gemmata 97 100 GQ250406_A_aff._pantherina 74 HQ604824_A_gemmata_ HM240517_A_pantherina 59 JF899545_A_gemmata EU071911_A_muscaria 100 AB080787_A_muscaria EU071960_A_muscaria KF651010_A_subfrostiana 100 60 JN943173_A_subfrostiana JN943172_A_subfrostiana KJ638273_A_frostiana 63 KP313580_A_frostiana 98 KP313579_A_frostiana KP313582_A_frostiana KP224322_A_populiphila GQ250409_A_velosa sect. Vaginatae 80 KM658285_A_pseudovaginata

0.02 Figure 8. Molecular phylogenetic analysis of Amanita brunneopantherina based on ITS sequences. The evolutionary history was inferred by the Maximum Likelihood method using Jukes-Cantor model. The analysis involved 41 nucleotide sequences. There were a total of 652 positions in the final dataset. Sequences generated from LAH35014 are marked with .

35

Amanita flavipes S. Imai, Bot. Mag., Tokyo 47: 428 (1933) (Figures 9 & 10)

Pileus 2.5–3.5 cm broad, convex to flat at maturity; cuticle bright yellow (10Y9/12) and / or golden brown (10YR7/10). Squamules white to light brown (7.5YR5/4); surface dry; margins rough, incurved and rimose at maturity. Lamellae off white (7.5Y9/2), up to 0.3 cm broad, adnexed, sub distant to close, edges entire. Lamellulae few, (2/3 to the length of lamellae). Stipe 5–9.5 cm × 0.6–1 cm, apex slightly wide, base bulbous, napiform, 1–2 cm in diameter, central, cylindrical, surface dry slightly fibrillose to floccose or scaly, bright yellow (10Y9/12) from the top becoming milky white or off white (7.5Y9/2) with light yellow (10Y9/8) tinge in patches. Sometimes scaly having brown (7.5Y?R5/8) dry scales. Brownish (5Y7/4) appearance in the form of rings below the annulus also observed. Annulus descending white to yellow (10Y9/12).

Basidiospores [30/6/6] (6.8) 8–9.3 (9.7) × (4.2) 5–6.3 (7.5) µm, Q = (1.3) 1.4–1.5 (1.6), avQ = 1.5 ± 0.01; ellipsoid, exine smooth, apiculus prominent, grayish to hyaline in 5% KOH. Basidia 8–9.3 × 3.33–42 µm, clavate having 2–4 starigmata, thin walled, wall up to 3 µm thick, hyaline 5% KOH, clamps absent at the base of basidia. Cystidia 6.6–8 × 23.3–26.3 µm, clavate, hyaline in 5% KOH. Pileipellis hyphae filamentous 1.7–5.5 µm in diameter, septa not observed, hyaline in KOH. Squamules hyphae globose to sub globose 17.2–20.2µm in diameter with a filament having fusiform terminals 1.6–6.5 µm in diameter, hyaline in 5% KOH. Stipitipellis hyphae 1.5–7 µm wide, filamentous, branched, broad fusiform terminals, hyaline in 5% KOH, septa not observed.

Material examined: Pakistan, Khyber Pakhtunkhwa province, Malakand division, Swat district, Kalam, 2400 m asl, on soil under Cedrus deodara, 4 Sep 2013, Sana Jabeen K3- 8; SJ8 (LAH35016); K2-21; SJ23 (LAH35017); Mashkun 2500 m asl, on soil under Cedrus deodara, 5 Sep 2013, Sana Jabeen K4-4; SJ1 (LAH35015); K4-18; SJ32 (LAH35018); K4-5; SJ42 (LAH35019); K4-30; SJ43 (LAH35020); K4-3; SJ49 (LAH35021); K4-18; SJ63 (LAH35022); K4-2; SJ64 (LAH35023).

Molecular phylogenetic characterization (Figure 11)

Sequences of the ITS region of Amanita flavipes consist of 716–805 nucleotide base pairs using ITS1F and ITS 4 as forward and reverse primers respectively. Consensus sequence of 619 base pairs from collection LAH35015 was BLAST searched at NCBI. It

36 showed 100% sequence similraity with A. flavipes (KF245911) from South Korea with 99% query cover and 0.0 E value.

For phylogenetic reconstruction, identical sequences from GenBank were retrieved. Taxa from section Phalloideae (Fr.) Quél. were chosen as out group. The sequences generated from local specimens clustered with A. flavipes from China (AY436455 & FJ375327), South Korea (KF245911) and Taiwan (KC185406) in section Validae (Fr.) Quél. of subgenus Lepidella (E.-J. Gilbert) Veselý with a very strong boot strap support.

Comments

Amanita flavipes is commonly known as "Asian yellow dust Amanita". It was originally described from Japan and widely distributed in Asian regions, particularly China, India, Japan, Pakistan, and South Korea (Ahmad et al., 1997; Zhang et al., 2004; Sanmee et al., 2008). The species ranges in color from yellow to yellowish brown, brownish orange, darker in the center and paler towards the margin, sometimes having grayish tints. Zhang et al. (2004) described a number of genetic clades within Amanita flavipes on the basis of molecular evidence supported by macroscopic differences.

37

Figure 9. Morphology of Amanita flavipes. A–H. Basidiomata. A. LAH35018; B & C. LAH35017; D. LAH35015; E. LAH35020; F. LAH35019; G. LAH35023; H. LAH35021. Bars: A = 2 cm; B & C = 1.5 cm; D = 1.2 cm; E =0.7 cm; F = 1 cm; G = 1.5 cm; H = 1.8 cm.

38

B A

C D

E F Figure 10. Anatomy of Amanita flavipes. A–F. LAH35016. A. Cystidia; B. Basidiospores; C. Basidia; D. Squamules; E. Pileipellis; F. Stipitipellis. Bars: A–F = 20 µm.

39

KJ638283_A_flavoconia KM373250_A_flavoconia GQ415317_A_flavoconia JN020966_A_flavoconia 100 JF313655_A_flavoconia KJ638281_A_flavoconia GQ415317_A_flavoconia JF313655_A_flavoconia 97 JF313657_A_flavoconia JN020967_A_flavoconia KJ638282_A_flavoconia KF937300_A_flavoconia 74 KF937299_A_flavoconia 100 KF937301_A_flavoconia FJ890029_A_flavoconia KR919762_A_morrisii KT213441_A_morrisii 100 KR919760_A_morrisii 17 KP284300_A_morrisii 58 KP284299_A_morrisii A flavipes SJ49 72 A flavipes SJ 64 A flavipes SJ43 A flavipes SJ42 KC185406_A_flavipes KF245911_A_flavipes 39 A flavipes SJ32 A flavipes SJ23 100 A flavipes SJ8 FJ375327_A_flavipes A flavipes SJ63 AY436455_A_flavipes A flavipes SJ1 AF085486_A_spissa JF907760_A_rubescens 100 AM084702_A_rubescens 74 63 AY436453_A_excelsa 99 KJ535438_A_novinupta KJ535437_A_novinupta 95 KF561974_A_novinupta GQ166902_A_flavorubescens sect. Validae JF313650_A_flavorubescens 65 90 EU819454_A_cf_flavorubesce AJ889923_A_rubescens FJ890031_A_rubescens JF313652_A_rubescens 81 AM087243_A_rubescens JF313652_A_rubescens 89 FJ890030_A_rubescens 71 JF313651_A_rubescens AJ889922_A_rubescens 47 93 KM409441_A_rubescens KF245919_A_rubescens 99 KF245918_A_rubescens 93 AB015682_A_rubescens 54 99 KM052535_A_rubescens 65 KM052530_A_rubescens JF273507_A_rubescens FJ890033_A_brunneoloculari 80 EU819464_A_rubescens 33 JN020972_A_rubescens GQ250403_A_novinupta 89 KC152065_A_novinupta 84 KC152067_A_novinupta 79 KC152066_A_novinupta 36 JX515562_A_franchetii 100 69 93 JX515561_A_franchetii 51 JX515563_A_franchetii AF085485_A_aspera AB0156951_A_sp 100 KJ466372_A_aff_fritillaria 39 KJ466373_A_aff_fritillaria KJ638284_A_rubescens 37 KF245910_A_spissa 54 EF493271_A_spissa 94 EF493270_A_spissa 97 AJ889924_A_spissa AJ889924_A_spissa AB015679_A_citrina AB015677_A_porphyria 100 100 JF899548_A_porphyria KC755034_A_phalloides EF493032_A_virosa sect. Phalloideae 82 KJ638292_A_bisporigera

0.05 Figure 11. Molecular phylogenetic analysis of Amanita flavipes based on ITS sequences. The evolutionary history was inferred by the Maximum Likelihood method using Jukes-Cantor model. The analysis involved 84 nucleotide sequences. There were a total of 724 positions in the final dataset. Sequences generated during present study are marked with .

40

Amanita glarea nom. prov. (Figure 12)

Etymology: The specific epithet refers to the gravel color of the pileus.

Pileus 3.8 cm broad, convex to flat at maturity; slightly umbonate, cuticle thin, striate, grayish brown (5YR4/2) from the middle becoming light gray (10Y7/2). Veil remnants not observed; surface dry; margins slightly incurved and rimose. Lamellae off white (7.5Y9/2), up to 0.3 cm broad, adnexed, sub distant to close, edges entire. Lamellulae few, (2/3 to the length of lamellae). Stipe 7.7 cm long, 0.5 cm from the apex becoming broad from the middle up to 0.8 cm, up to 1.5 cm with a prominent sac like volva, central, cylindrical, surface dry, granular from the top, becoming smooth towards the base, brown (2.5YR3/4) to off white (7.5Y9/2). Annulus absent.

Basidiospores [20/1/1] (10) 10.1–11.7 (12) × (9.6) 10.4–11 (11.5) µm, Q = 0.97– 0.96, avQ = 0.98 ± 0.1; non amyloid, globose, surface smooth, hyaline to grayish in 5% KOH. Basidia 32–36 × 11.2–14.4 µm, clavate having 2 and 4 starigmata, thin walled, wall up to 3 µm thick, hyaline 5% KOH. Cystidia 11.2–12 × 33.6–44 µm, clavate, hyaline in 5% KOH. Pileipellis hyphae filamentous, 11.3–21.6 µm in diameter, infrequently septate, hyaline in KOH. Squamules absent. Stipitipellis hyphae filaments, 9.3–24.7 µm wide, branched, hyaline in 5% KOH, septa infrequent.

Material examined: Pakistan, Khyber Pakhtunkhwa province, Malakand division, Swat district, Mashkun, 2500 m asl, on soil under Cedrus deodara, 5 Sep 2013, Sana Jabeen & Abdul Nasir Khalid K4-34; SJ44 (LAH35044).

Molecular phylogenetic characterization (Figure 13 & 14)

Sequences of the 610–616 nucleotide base pairs obtained from ITS products using ITSIF and ITS4 primers were trimmed at conserve motifs. A sequence of 512 nucleotides was BLAST search at NCBI. It was found 99% identical to unidentified Amanita spp. (KT006761 & KP258996) from Canada, Malawi (JF710841), USA (KP711843) and Zambia (JF710842).

Sequencing of the PCR products of LSU yielded upto 588 base pairs by using LR0R and LR5 primers. BLAST searched at NCBI showed 99% identity to sequences from Malawi (JF710817) and China (AF024442) with 94–96% query cover and 0.0 E value.

41

Identical sequences of ITS and LSU regions were retrieved from GenBank to construct phylogenetic tree. Taxa from section Caesareae Singer ex Singer were chosen as out group. The sequences generated from the Pakistani collection clustered with the taxa from Canada, France, Malawi, Russia USA and Zambia in same clade within section Phalloideae (Fr.) Quél. of subgenus Amanita Pers.

Comments

Amanita glarea is treated as a new taxon. It is characterized by its grayish brown, thin pileal surface, with striations, brown to off white stipe having a sac like volva. Anatomically it is characterized by its globose basidiospores. Molecular data based on ITS region showed that the sequences from Pakistan are 97.5–98.4% similar to other unidentified taxa in the same clade with 1–2% genetic divergence. Taxa in the section Vaginatae are more or less similar morphologically. Morphologically the species is closely related to A. vaginata (Bull.) Lam. with no remarkable differences accept the color of the pileus which is more brownish as compared to A. glarea. It is also comparable with A. velosa (Peck) Lloyd, a close relative in the same clade. Amanita glarea differs from A. velosa by the lack of umbo on its pileus. Moreover, A. velosa bears pale orange to pale salmon color when young that becomes more brownish (Jenkins, 1986; Tulloss, 2005). Amanita glarea did not show any orangish coloration.

42

Figure 12. Morphology and anatomy of Amanita glarea. A–F. LAH35044 (holotype). A & B. Basidiomata; C. Basidiospores; D. Basidia and Cystidia; E. Pileipellis F. Stipitipellis. Bars: A = 1.2 cm; B = 1.3 cm; C = 6 µm; D = 8.8 µm; E & F = 24 µm.

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98 KP258995_A_supravolvata 73 KM658285_A_pseudovaginata A glarea SJ44 92 A glarea SJ44b 92 KT006761_A_sp_subnigra KP711843_A_sp_subnigra KP258996_A_sp_subnigra 42 JF710842_A_sp_Kew JF710841_A_sp_Kew 99 KM658303_A_aff_vaginata KM658302_A_aff_vaginata 96 KM658299_A_vaginata 63 KM576299_A_sp 66 KP711841_A_sp 99 JQ976014_A_sp KJ638266_A_crocea AY918961_A_velosa 89 GQ250409_A_velosa 22 100 DQ974692_A_velosa 55 EU909453_A_velosa 96 KM658296_A_aff_vaginata 84 KJ638267_A_sp KM658298_A_vaginata KP224316_A_sp_albiceps KP224327_A_protecta 59 100 KP224328_A_protecta 48 KP224324_A_protecta KP224326_A_protecta 65 89 KP224322_A_populiphila KP224318_A_populiphila sect. Vaginatae 97 KP224320_A_populiphila 44 75 KP224321_A_populiphila JQ912665_A_crocea JF907762_A_oblongispora 99 JF907758_A_beckeri 92 JF907758_Amanita_beckeri 91 KF017948_A_vaginata KF017948.1|_Amanita_vaginata 35 69 KF017949_Amanita_vaginata KP004955_A_vaginata 99 83 KF017949_A_vaginata 71 JN182880_Amanita_lignitincta 91 KP348228_A_lignitincta 99 JN182881_A_constricta JN182880_A_lignitincta 100 JQ347048_A_lignitincta 32 FJ441045_Amanita_lignitincta 66 KM658297_A_aff_lignitincta KM658297_Amanita_aff_lignitincta 99 KM595044_A_aff_lignitincta 86 86 KM658292_Amanita_aff_lignitincta KP662537_A_sp_rhacopus KP224339_A_sp_rhacopus KP224338_A_sp_rhacopus KP224337_A_sp_rhacopus 99 KF359589_A_cf_constricta KJ638264_A_sp KF007934_A_constricta GU220373_A_cf_constricta JQ512095_Amanita_loosii sect. Caesareae

0.05 Figure 13. Molecular phylogenetic analysis of Amanita glarea based on ITS sequences. The evolutionary history was inferred by the Maximum Likelihood method using Jukes-Cantor model. The analysis involved 60 nucleotide sequences. There were a total of 676 positions in the final dataset. Sequences generated from LAH35044 are marked with .

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52 FJ705276 Amanita submembranacea 88 KU867877 Amanita olivaceogrisea HQ539739 Amanita sinicoflava 92 98 KT317710 Amanita mortenii HQ539684 Amanita constricta AY228351 Amanita constricta 48 80 90 DQ384583 Amanita constricta 100 KU139437 KF021668 Amanita ceciliae 22 100 KF021669 Amanita ceciliae 92 KU139438 Amanita ceciliae KF021680 Amanita orientifulva 100 LC09876 Amanita orientifulva LC098755 Amanita orientifulva KT779068 Amanita fulva 78 32 KU139446 Amanita fulva 96 KU139448 Amanita fulva LC098759 Amanita fulva KP866162 Amanita sp daimonioctan 28 KP284297 Amanita sp daimonioctan KP284296 Amanita sp daimonioctan KP284298 Amanita sp daimonioctan EU522724 Amanita aff vaginata GQ250425 Amanita sp sect. Vaginatae 98 KM658309 Amanita battarrae 24 KM658305 Amanita battarrae 78 AF024481 Amanita umbrinolutea KU248121 Amanita friabilis 100 KU248120 Amanita friabilis KU248119 Amanita friabilis HQ539695 Amanita friabilis KU186821 Amanita sp rhacopus 74 KP221313 Amanita sp rhacopus 44 KU186820 Amanita sp rhacopus KP866170 Amanita sp texasorora KP662534 Amanita sp texasorora 80 KP662533 Amanita sp texasorora Amanita glarea SJ44a 78 Amanita glarea SJ44b JF710817 Amanita sp AF024442 Amanita brunneofuligine 98 KM658313 Amanita aff vaginata 54 KM658312 Amanita aff vaginata KP224343 Amanita populiphila 58 KP221304 Amanita populiphila 100 KP224344 Amanita populiphila KP221314 Amanita populiphila JQ512080 Amanita loosii sect. Caesareae

0.01 Figure 14. Molecular phylogenetic analysis of Amanita glarea based on LSU sequences. The evolutionary history was inferred by the Maximum Likelihood method using Jukes-Cantor model. The analysis involved 48 nucleotide sequences. There were a total of 567 positions in the final dataset. Sequences generated from LAH35044 are marked with .

45

Amanita swatica nom.prov. (Figures 15 & 16)

Etymology: The specific epithet refers to the district name from where the samples were collected.

Pileus upto 6 cm broad, convex to plano convex at maturity; cuticle reddish brown (10R2/6) from the centre, dark brown (10R3/6) to pale yellow (5Y8/6) towards the margins, sometimes dark brown color is present in the form of a network on light yellow base, surface dry. Veil remnants gray (5Y7/2), common, sometimes absent, surface dry; margins smooth, incurved when young becoming flat and rimose at maturity. Lamellae off white (2.5BG4/2) to cream (5Y9/4) up to 0.3 cm broad, adnexed, subdistant to close, margins entire. Lamellulae diverse in size, three tiered. Stipe 10 cm × 1.3 cm, base up to 2 cm wide becoming narrower towards the apex, central, cylindrical, surface smooth grainy from the apex becoming fibrillose towards the base, white cottony patches observed towards the apex on brown (10R6/12) context, brown striations are present towards the base, surface dry, sometimes scaly, becoming pinkish on bruising. Annulus prominent, present near the apex, skirty, papery, off white towards the gills and brownish from the opposite side.

Basidiospores [50/2/2] 7.1–9.6 × 6.2–7.0 µm, Q = 1.1–1.3, avQ = 1.17, globose to subglobose, guttulate, exine smooth, apiculus prominent, grayish in 5% KOH. Basidia 232–255 × 47–55.5 µm, guttulate, clavate, having 2–4 starigmata, thin walled, wall up to 3 µm thick, hyaline 5% KOH. Cystidia 80–178 × 32–49 µm, clavate, hyaline in 5% KOH. Pileipellis 2.7–40 µm, clavate terminals, hyaline in 5% KOH. Squamules hyphae globose to elliptical or fusoid 20–52 × 29–70 um with a filament 2.2–5.6 µm, brownish in 5% KOH. Stipitipellis hyphae 2.2–13.6 µm, clavate to cylindrical, frequently septate, hyaline in 5% KOH.

Material examined: Pakistan, Khyber Pakhtunkhwa province, Malakand division, Swat district, Mashkun, 2500 m asl, on soil under Cedrus deodara, 5 Sep 2013, Sana Jabeen K4-1; SJ33 (LAH35025); K4-27; SJ88 (LAH35026); 1K4; SJ67 (LAH35207).

Molecular phylogenetic characterization (Figure 17)

Sequencing of the PCR products of ITS region of Amanita swatica produced fragments of 724–784 base pairs by using ITS1F and ITS4 primers. Consensus sequence of 619 base pairs was obtained by trimming at conserved sites. BLAST searched at NCBI

46 revealed that A. swatica is 99% identical to sequences from Denmark (AJ889923 & AM087243), Russia (JF313651) and USA (JF313652) with 99% query cover and 0.0 E value.

For phylogenetic analysis, closest sequences from the GenBank were retrieved . Taxa from section Phalloideae (Fr.) Quél were added in the final alignment file as outgroup. The sequences generated during this study clustered in section Validae (Fr.) Quél, of subgenus Lepidella. Amanita swatica clustered in a clade with A. novinupta Tulloss & J. Lindgr. and A. rubescens Pers. It get separated from A. rubescens forming a sister clade with 73% bootstrap value.

Comments

Amanita swatica nom. prov. is characterized by its reddish brown to dark brown pileus surface with some network of brownish coloration on a pale yellow context and globose to subglobose basidiospores. In comparison with A. rubescens, pileus color of A. rubescens is brown to yellowish brown. It differs from A. swatica where the pileus is reddish brown from the centre becoming brown forming a network on the pale yellow context towards the margins. No such coloration pattern has been found in A. rubescens. Anatomically, A. swatica also differs from A. rubesence on the basis of size. The spore size also differs, basidiospores of A. rubescens are comparatively larger (7.3–9.5 × 5.9–7.0) and rarely globose than that of A. swatica (Kuo, 2013). Amanita novinupta, another closely related taxon, is morphologically distinct from A. swatica by its white to pinkish pilues color becoming darker with age and elliptical spores (Tulloss & Lindgren, 1994). Molecular data analysis also placed this taxon in separate lineage.

47

Figure 15. Morphology of Amanita swatica. A–D. Basidiomata. A & B. LAH35025 (holotype); C & D. LAH35027. Bars: A = 2.25 cm; B = 1.75 cm; C & D = 1.5cm.

48

Figure 16. Anatomy of Amanita swatica. A–F. LAH35025. A. Basidiospores; B. Pileipellis; C. Cystidia; D. Basidia. E. Stipitipellis; F. Squamules hyphae. Bars: A = 5 µm; B-E = 10 µm; F = 20 µm.

49

JF313652_A_rubescens AJ889923_A_rubescens AM087243_A_rubescens 80 JF313652_A_rubescens FJ890030_A_rubescens FJ890031_A_rubescens 73 JF313651_A_rubescens AJ889922_A_rubescens 89 KM409441_A_rubescens A swatica SJ88 A swatica SJF33 69 58 72 A swatica SJ33b JF273507_A_rubescens 75 KM052535_A_rubescens 100 AB015682_A_rubescens 64 KM052530_A_rubescens FJ890033_A_brunneoloculari 97 EU819464_A_rubescens 90 JN020972_A_rubescens GQ250403_A_novinupta 96 KC152065_A_novinupta 87 KC152067_A_novinupta 81 KC152066_A_novinupta KJ535438_A_novinupta 99 KF561974_A_novinupta KJ535437_A_novinupta 70 95 GQ166902_A_flavorubescens JF313650_A_flavorubescens 68 92 EU819454_A_cf_flavorubesce AM084702_A_rubescens 61 AY436453_A_excelsa 100 JF907760_A_rubescens AF085486_A_spissa AY436455_A_flavipes 100 KF245911_A_flavipes FJ375327_A_flavipes 52 KC185406_A_flavipes sect. Validae KP284300_A_morrisii KP284299_A_morrisii 100 KR919760_A_morrisii 43 KT213441_A_morrisii KR919762_A_morrisii KF937300_A_flavoconia 100 KF937299_A_flavoconia 74 46 KF937301_A_flavoconia FJ890029_A_flavoconia JN020967_A_flavoconia GQ415317_A_flavoconia 99 JN020966_A_flavoconia JF313655_A_flavoconia KJ638281_A_flavoconia GQ415317_A_flavoconia 93 JF313655_A_flavoconia JF313657_A_flavoconia KM373250_A_flavoconia KJ638283_A_flavoconia KJ638282_A_flavoconia JX515562_A_franchetii 74 JX515561_A_franchetii 98 JX515563_A_franchetii 66 AF085485_A_aspera AB0156951_A_sp 99 KJ466372_A_aff_fritillaria 42 KJ466373_A_aff_fritillaria KJ638284_A_rubescens KF245910_A_spissa 42 EF493271_A_spissa 94 EF493270_A_spissa 99 AJ889924_A_spissa AJ889924_A_spissa AB015679_A_citrina AB015677_A_porphyria 100 100 JF899548_A_porphyria EF493032_A_virosa KC755034_A_phalloides sect. Phalloideae 100 78 KJ638292_A_bisporigera

0.05 Figure 17. Molecular phylogenetic analysis of Amanita swatica based on ITS sequences. The evolutionary history was inferred by the Maximum Likelihood method using Jukes-Cantor model. The analysis involved 76 nucleotide sequences. There were a total of 711 positions in the final dataset. Sequences generated during this study are marked with .

50

Genus Boletus

Boletus himalayensis nom. prov. (Figures 18 & 19)

Etymology: The specific epithet refers to Pakistan's part of Himalayan ranges, from where the sample was first collected.

Pileus up to 6.5 cm wide, convex to planoconvex; surface dry, smooth, ruptured, whitish context visible between cracks, yellowish brown (10YR7/8) to dark brown (10R6/16), margins entire, rimose, incurved to straight. Pore surface free and approximate, 11–18 mm deep, off white to yellowish brown (2.5Y7/6), pores circular, up to 3/mm. Stipe up to 4.7 × 2.4 cm, base and apex narrow up to 1 cm, central, surface densely reticulate, off white reticulations on yellowish to brownish context at the upper 1/3 of the stipe, basal part off white, non reticulated.

Basidiospores [50/2/1] 25–37 × 10–14 µm, Q = 2.1–3.3, avQ = 2.3, ellipsoid, exine smooth, apiculus prominent, Basidia 21–32 × 12–14 µm, clavate 4-spored, thin walled, guttulate, hyaline 5% KOH. Cystidia, long, 21.5–31 × 6–8 µm, densely clustered at the edges of tubes, clavate to subclavate or cylindrical, hyaline in 5% KOH, . Pileipellis cellular, cells elongated, 9–15 µm wide, terminals clavate, hyaline in 5% KOH. Stipitipellis hyphae thin, 7–12 µm wide, filamentous, branched, septa not observed, hyaline in 5% KOH.

Material examined: Pakistan, Khyber Pakhtunkhwa province, Malakand division, Swat district, Mashkun, 2500 m asl, on soil under Cedrus deodara, 5 Sep 2013, Sana Jabeen & Abdul Nasir Khalid MT2WB; SJ6 (LAH35028); MT2D; SJ7 (LAH35029).

Molecular phylogenetic characterization (Figures 20 & 21)

Sequencing of the PCR products of ITS region of Boletus himalayensis consisted of 790–811 base pairs by using ITS1F and ITS4 primers. Consensus sequence of 698 and 701 base pairs was obtained by trimming the motifs at conserve sites. Sequence from LAH35028 collection was BLAST searched at NCBI. It showed 99% identity to Boletus reticuloceps (Zang et al.) Wang & Yao sequences from China (EU231968, JN563885, JN563882, FJ548566, JN563887 & JN563886) with 89–99% query cover and 0.0 E value.

51

Sequencing of the PCR products of LSU yielded 936–948 base pairs by using LR0R and LR5 primers. A sequence of 948 base pairs from LR0R primer was BLAST searched at NCBI. It showed 98% identity to Boletus fibrillosus Thiers (KF030343 & KF030344) from USA, and B. reticuloceps (JN563843 & KF112454) from China with 88–92% query cover. It was also found 97% identical to B. edulis Bull. sequences (DQ071747 & HQ161848) from Germany and USA with 94 and 92% query cover respectively.

To study phylogeny, closely related ITS and LSU sequences were retrieved from the GenBank. For ITS B. edulis (GU198977) and for LSU tree, Porphyrellus brunneus McNabb (JX889646) were chosen as out group. The sequences generated from ITS clustered with similar taxa from China forming a clade of their own with 73% bootstrap value. Sequences from LSU region clustered in a same clade with Chinese taxa but formed their own lineage with 82% bootstrap value.

Comments

Boletus himalayensis nom. prov. clustered with similar taxa in the section Boletus which is characterized by reticulated stipe, at least in apical part, whitish context with no color change upon bruising and whitish pore surface in young stages (Thiers, 1975; Singer, 1986). Boletus himalayensis is characterized by smooth to cracked pileus surface and whitish to yellowish pore surface which differentiates its closest relative B. retciculoceps which has velvety, wrinkled to squamules pileus surface. Anatomically, B. himalayensis can be differentiated due to sterile tube edges having long dense cluster of cystidioid elements. Molecular phylogenetic analysis based on ITS and LSU also support the identification of B. himalayensis as a new taxon.

52

A

B C Figure 18. Morphology of Boletus himalayensis. A–C. Basidiomata. A. LAH35028 (holotype); B & C. LAH35029. Bars: A = 1.5 cm; B & C = 1.2 cm.

53

A

B C

D E

Figure 19. Anatomy of Boletus himalayensis. A–E. LAH35029. A. Basidiospores; B. Basidia and basidiole; C. Cystidia; D. Stipitipellis; E. Pileipellis. Bars: A = 13 µm; B & C = 5 µm; D = 22 µm; E = 19 µm.

54

KJ131224 B reticuloceps JN563884 B reticuloceps KJ828812 B reticuloceps 73 KJ131226 B reticulocepshimalayensis SJ7 SJ7 48 KJ131225 B reticulocepshimalayensis SJ6 SJ6 JN563882 B reticuloceps FJ378737 Uncultured Boletus 63 FJ378736 Uncultured Boletus JN563883 B reticuloceps JN563887 B reticuloceps JN563886 B reticuloceps 67 JN563885 B reticuloceps GU198977 B edulis

0.005 Figure 20. Molecular phylogenetic analysis of Boletus himalayensis based on ITS sequences. The evolutionary history was inferred by the Maximum Likelihood method using General Time Reversible model. The analysis involved 13 nucleotide sequences. There were a total of 711 positions in the final dataset. Sequences generated from during this study are marked with .

55

DQ071747_Boletus_edulis AF291300_Boletus_edulis 65 AF462352_Boletus_edulis AF462353_Boletus_edulis AF462354_Boletus_edulis KF112455_Boletus_edulis 93 KC184482_Boletus_edulis_var_grandedulis 65 KC900423_Boletus_edulis_var_grandedulis KC900411_Boletus_rubriceps KC900407_Boletus_rubriceps KC900408_Boletus_rubriceps 34 47 KC900412_Boletus_rubriceps 100 KF030342_Boletus_aurantioruber KF030341_Boletus_subcaerulescens AF462358_Boletus_pinophilus KF030340_Boletus_subalpinus 98 AF462359_Boletus_pinophilus 64 DQ273495_Uncultured_Boletaceae AF071457_Boletus_edulis 79 KC184485_Boletus_regineus KF030339_Boletus_aereus KF030343_Boletus_fibrillosus KF030344_Boletus_fibrillosus 93 Boletus himalayensis SJ6b Boletus himalayensis SJ6 82 JN563843_Boletus_reticuloceps 68 KF112454_Boletus_reticuloceps 45 JN563847_Boletus_sp AF462355_Boletus_cf_edulis

100 KC184480_Boletus_barrowsii JQ172787_Boletus_sp 36 JN563852_Boletus_sp 49 JN563851_Boletus_sp JX889646_Porphyrellus_brunneus

0.01 Figure 21. Molecular phylogenetic analysis of Boletus himalayensis based on LSU sequences. The evolutionary history was inferred by the Maximum Likelihood method using General Time Reversible model. The analysis involved 34 nucleotide sequences. There were a total of 622 positions in the final dataset. Sequences generated from LAH35028 are marked with .

56

Genus Hortiboletus

Hortiboletus rubellus (Krombh.) Simonini, Vizzini & Gelardi, 244: 1 (2015) (Figure 22)

Pileus 6.5 cm wide, flat, margins uplifted, surface dry bearing cracks, pinkish brown (7.5R5/14) to brown (10R5/14). Context yellowish, slightly bluing upon exposure. Stipe 7.5 × 1 cm, narrower from the centre, base 1.5 cm wide, apex 1.4 cm wide, central, solid, pinkish brown striations becoming reddish (7.5R4/14) towards the apex, apex and base pale yellow (2.5Y8/8). Hymenium yellowish (10YR7/10), adnate, slightly turns blue upon bruising, pores 2–3/mm, tubes up to 9 mm deep.

Basidispores fusiform, apiculate, smooth, 13–15 × 5–7 µm. Basidia long, clavate, guttulate, sterigmata 2–4, long, 47–54 × 9–15 µm. Cystidia cylindrical, clavate to subclavate, 44–49 × 10–11.5 µm.

Material examined: Pakistan, Khyber Pakhtunkhwa province, Malakand division, Swat district, Mashkun, 2500 m asl, on soil under Cedrus deodara, 5 Sep 2013, Sana Jabeen & Abdul Nasir Khalid SJ113 (LAH35030).

Molecular phylogenetic characterization (Figure 23)

Sequencing of the PCR products of ITS region of Hortiboletus rubellus yielded fragments of 863 and 876 base pairs by using ITS1F and ITS4 primers. Consensus sequence of 780 base pairs was obtained by trimming the motif and BLAST searched at NCBI. It showed 99% identity to sequences from Pakistan (KJ802929 & KJ802928), Austria (EF644119), France (KM576320) and Montenegro (JQ685725) with 89–98% query cover and 0.0 E value.

For phylogenetic tree, ITS sequences from closely related taxa were retrieved from the GenBank. Porphyrellus brunneus McNabb (JX889646) was added in the final alignment file to root the tree. The sequence generated from LAH35030 was clustered with the similar taxa from Austria, France, Montenegro and Pakistan with 98% boot strap value.

57

Comments

Hortiboletus Simonini, Vizzini & Gelardi is a newly described genus (Vizzini, 2015). Hortiboletus rubellus is the specimen of this genus and commonly known as “ruby bolete”. It was named as Boletus rubellus Krombh. in 1836. Another commonly used binomial name Boletus versicolor (Rostk.) was used for this taxon which later reduced to synonymy. Its present specific epithet rubellus is Latin for "somewhat red" (Simpson, 1979). Boletus rubellus, one of the pored basidiomycetes to be placed in the genus Quél. as Xerocomus rubellus (Krombh.) Quel. in 1896 (Lamaison & Polese, 2005). Later, the members of this genus were placed in Xerocomellus Sutara (Nuhn et al., 2013). The was transferred Hortiboletus in 2015, based on molecular evidence indicating its genetic dissimilarity to Boletus and Xerocomellus due to smooth spores in all Hortiboletus species.

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A B

C D E

Figure 22. Morphology and anatomy of Hortiboletus rubellus. A–E. LAH35030. A & B. Basidiomata; C. Basidiospores; D. Basidia; E. Cystidia. A & B = 6.5 cm; C, D & E = 10 µm.

59

JX030290_Boletus_aff_rubellus 69 JX030290_Boletus_aff_rubellus 72 JX030209_Boletus_aff_rubellus

55 GQ166883_Boletus_rubellus GQ166876_Boletus_rubellus 87 44 GQ166879_Boletus_campestris 67 GQ166888_Boletus_rubellus 29 EU819452_Boletus_fraternus

KT319647_Hortiboletus_sp 20

46 Hortiboletus rubellus SJ113 KJ802929_Boletus_rubellus

98 KJ802928_Boletus_rubellus 37 EF644119_Xerocomus_rubellus 39

49 KM576320_Boletus_sp 66 JQ685725_Xerocomus_rubellus

KJ609174_Xerocomus_sp

EU819460_Boletus_rubellus

99 JX434687_Boletus_cf_rubellus 100 AB211279_Boletus_cf_rubellus

JQ003627_Phylloporus_pumilus

0.05

Figure 23. Molecular phylogenetic analysis of Hortiboletus rubellus based on ITS sequences. The evolutonary history was inferred by the Maximum Likelihood method using Jukes-Cantor model. The analysis involved 20 nucleotide sequences. There were a total of 989 positions in the final dataset. Sequences generated from LAH35030 are marked with .

60

Genus Neoboletus

Neoboletus luridiformis (Rostk.) Gelardi, Simonini & Vizzini, Index Fungorum, 192: 1 (2014) (Figure 24)

Pileus up to 8 cm wide, covex becoming plane at maturity, surface dry, smooth, gray (5Y5/2) to brown (10YR7/6) or grayish brown (10YR4/4), sometimes cracks are present and off white (5Y9/4) context visible from within, margins incurved, entire smooth. Stipe up to 9 cm long, thick up to 4 cm, solid, centric, dry, pinkish (6R6/14) on an off white context, apex becoming yellow (2.5Y8/12) towards the hymenium. Pore surface reddish (7.5R5/16), yellowish (2.5Y8/12) from the inside, adnate, color change to blue upon bruising, pores frequent, up to 1.7cm deep.

Basidispores [40/4/4] 13–18 × 5–8 µm, ellipsoid, thick walled, smooth. Basidia 20–27 × 9–13 µm, clavate, 2–4 sterigmata, hyaline in 5% KOH. guttulate. Cystidia 30–48 × 10–13 µm subclavate to cylindrical, hyaline in 5% KOH.

Material examined: Pakistan, Khyber Pakhtunkhwa province, Malakand division, Swat district, Kalam, 2400 m asl, on soil under Cedrus deodara, 3 Sep 2013, Sana Jabeen K2-3 (LAH35032); K2-4 (LAH35033); Mashkun, 2500 m asl, on soil under C. deodara, 5 Sep 2013, Sana Jabeen K411MT; SJ4 (LAH35031; GenBank KM199730); MTT (LAH35034); MT (LAH35035).

Molecular phylogenetic characterization (Figure 25)

Sequencing of the PCR products of ITS region of Neoboletus luridiformis yielded 791–850 base pairs by using ITS1F and ITS4 as forward and reverse primers. A consensus sequence of 695 base pairs was obtained by trimming the motif at conserve sites and BLAST searched at NCBI. It showed 99% genetic identity to Boletus erythropus Pers. sequences from Italy (DQ131633 & DQ131634) with 100% query cover and B. luridiformis Rostk. (KJ802930, KJ802931 & KJ802932) from Pakistan with 45–60% query cover.

For phylogenetic analysis, sequences derived from closely related taxa from GenBank were retrieved. Leccinum albellum (Peck) Singer (KM248926) was chosen as out group. The sequence generated from collection LAH35031 clustered in same clade

61 with Boletus erythropus and B. luridiformis from Italy and Pakistan with 71% boot strap value.

Comments

Neoboletus luridiformis, was known as Boletus luridiformis until 2014. It is commonly known as the "dotted stem bolete". Boletus luridiformis was originally described by Christian Hendrik Persoon in 1796 as B. erythropus, a name since reduced to synonymy (Phillips, 2006). The specific epithet was derived from the Greek meaning "red foot" referring to its red colored stalk (Liddell & Scott, 1980). Genetic analysis published in 2013 showed that B. luridiformis and many other red-pored boletes were well-removed from the Boletus and placed in a new genus characterized by the brown or reddish brown hymenophoral surface, the lack of reticulations on the stipe and an quick blue stain when bruised (Nuhn et al., 2013). Neoboletus luridiformis became the type species of the Neoboletus G.Wu & Zhu L.Yang in 2014 (Gelardi et al., 2014).

62

E F G

A B

C

Figure 24. Morphology and anatomy of Neoboletus luridiformis. A–D. Basidiomata. E–G. LAH35034. A–C. LAH35034; D LAH35035. E. basidiospores; F. cystidia; G. basidia. Bars: A–D = 2 cm; E = 5 µm, F = 14 µm; G = 6 µm.

63

HM347643 Boletus erythropus

61 KM576321 Boletus sp KM198319 Boletus pseudosulphureu KM198315 Boletus erythropus 64 HM347644 Boletus erythropus KM198314 Boletus erythropus

62 KM198318 Boletus erythropus DQ131634 Boletus erythropus KJ802930 Boletus luridiformis DQ131633 Boletus erythropus 71 KJ802931 Boletus luridiformis KJ802932 Boletus luridiformis 61 KM199730 Neoboletus luridiformis SJ4 KM198313 Boletus xanthopus KM198312 Boletus xanthopus 99 KM198311 Boletus xanthopus 34 KM198320 Boletus xanthopus KM027387 Boletus xanthopus 87 KM198317 Boletus xanthopus KM248926 Leccinum albellum

0.005 Figure 25. Molecular phylogenetic analysis of Neoboletus luridiformis based on ITS sequences. The evolutionary history was inferred by the Maximum Likelihood method using Jukes-Cantor model. The analysis involved 20 nucleotide sequences. There were a total of 707 positions in the final dataset. Sequences generated from LAH35031 are marked with .

64

Genus Xerocomellus

Xerocomellus rimosus nom. prov. (Figures 26 & 27)

Etymology: The specific epithet refers to cracked surface of the pileus.

Pileus up to 6.7 cm, convex to plano convex, spherical to irregular, surface dry, rough, light brown (10YR7/6) to dark brown (5YR3/6) or blackish brown (10YR2/4), abundant cracks are present, off white (5Y9/4) to light brown (10YR7/6) context visible from within, margins incurved, entire smooth and irregular. Stipe up to 7 cm long, thick up to 1.4 cm, base slightly narrow up to 1 cm, curved, solid, centric, dry, reddish (7.5R516) from the base, becoming yellow (5Y8/10) towards the apex. apex red (7.5R5/16) in the form of ring just below the attachment to the pileus, very apex yellowish (5Y8/10), context off white. Hymenium yellowish (5Y8/10 to brown (10YR6/8), becoming bluish upon bruising, pores frequent, up to 0.7cm deep.

Basidispores [30/2/2] 12.5–14.1 × 5.5–8.2 µm, ellipsoid, smooth, epiculus prominant. Basidia 17.4–30.2 × 7.1–9.5 µm, clavate, 1–4 sterigmate, thin walled, guttulate, contents visible. Cystidia 27.2–38.4 × 8.4–10.3 µm, cylindrical to subclavate, thin walled, hyaline in 5% KOH. Pileipellis hyphae 2.3 × 9.1 µm wide, long, filamentous, septate, hyaline in 5% KOH. Stipitipellis hyphae 4.3 × 14.7 µm wide, long, filamentous, frequently septate, hyaline in 5% KOH.

Material examined: Pakistan, Khyber Pakhtunkhwa province, Malakand division, Swat district, Kalam, 2400 m asl, on soil under Cedrus deodara 3 Sep 2013, Sana Jabeen K18 (LAH35037); Mashkun, 2500 m asl, on soil under C. deodara, 5 Sep 2013, Sana Jabeen K4-30; SJ65 (LAH35036).

Molecular phylogenetic characterization (Figures 28 & 29)

Sequencing of the PCR products of ITS region of Xerocomellus rimosus yielded 602–837 base pairs by using ITS1F and ITS4 primers. Consensus sequence of 670 base pairs was obtained by trimming the motif and BLAST searched at NCBI. It showed 93% identity to Xerocomus chrysenteron (Bull.) Quél. sequences from Germany (HQ207691– HQ207694), Scotland (JQ888153) and Switzerland (AF402139) with 99% query cover and 0.0 E value.

65

Sequencing of the PCR products of LSU yielded 931–1196 base pairs by using LR0R and LR5 primers. A sequence of 848 base pairs was BLAST searched at NCBI. It showed 99% genetic identity to Xerocomus chrysenteron. sequences from Ausria (AF514808 & AF514809), Sweden (AF347103) and Switzerland (AF402139) with 99– 100% query cover and 0.0 E value.

To reconstruct phylogeny, closely related ITS and LSU sequences were retrieved from the GenBank. For phylogenetic tree based on ITS, Phylloporus pumilus M. A. Neves & Hallin (JQ003627) while for LSU Tylopilus balloui (Peck) Singer (HQ161873) were chosen as out groups. The sequences generated during this study got separated from Xerocomellus chrysenteron and other taxa with a boot strap of 50% in ITS and 90% in LSU sequences based phylogram.

Comments

Xerocomellus rimosus nom. prov. is characterized by its brownish, dry, rough, rimose surface with pale brown context visible from within the craks and stipe reddish from the base becomming yellowish towards the stipe having a ring like pattern of red coloration just below the apex. Morphologically it is comparable with its sister taxon, X. chrysenteron. Xerocomelus chrysenteron bears cracks in the mature cap revealing a thin layer of light red flesh below the pileipellis (Laessoe, 1998), while in X. rimosus, red layer is absent, instead a pale brown context is visible. Moreover the stipe surface of X. chrysenteron bears reddish fibrils more or less on the whole stipe, but in X. rimosus, the base of the stipe is reddish, becoming yellowish towards the apex and apex bears a prominent red coloration in the form of ring just below the pileus attachment. Molecular phylogenetic analysis also support it a distinct species from X. chrysenteron based on ITS and LSU sequences.

66

Figure 26. Morphology of Xerocomellus rimosus. A–D. Basidiomata. A & B. LAH35037 (holotype); C & D. LAH35036; D. Bars: A & B = 0.8 cm; C & D = 1.2 cm.

67

Figure 27. Anatomy of Xerocomellus rimosus. A–E. LAH35036. A. Basidiospores; B. Cystidia; C. Basidia; D. Stipitipellis; E. Pileipellis. Bars: A–E = 10 µm.

68

KM213655 Xerocomellus sp 76 KM213661 Xerocomellus sp

92 KM213653 Xerocomellus sp AY372284 Xerocomus sp 86 95 AY372288 Xerocomus sp 45 KM213635 Xerocomellus sp 97 KM213636 Xerocomellus sp KM213665 Xerocomus truncatus 100 JX030249 sp 92 KJ882290 Hymenogaster behrii 94 KJ882288 Hymenogaster behrii KJ882289 Hymenogaster macmurphyi 82 KM213662 75575 Boletus rainisii AY372283 Boletus dryophilus A 54 73 AY372286 Boletus dryophilus 98 KM213646 Xerocomellus sp 99 HM190086 Xerocomus porosporus KM085410 Xerocomellus porosporus Xerocomellus rimosus SJ65 68 JF908799 Xerocomellus chrysentereron HQ207694 Xerocomus chrysenteron

50 JQ888153 Boletus chrysenteron HQ207693 Xerocomus chrysenteron AF402139 Xerocomus chrysenteron 99 EU350581 Xerocomus chrysenteron HQ207691 Xerocomus chrysenteron 71 HQ207692 Xerocomus chrysenteron KC581337 Xerocomellus zelleri 93 HM190092 Xerocomus pruinatus 100 HM190099 Xerocomus pruinatus 97 HM190088 Xerocomus pruinatus

71 KM213641 Xerocomellus sp KM213637 Xerocomellus sp 90 KM213640 Xerocomellus sp 87 KM213638 Xerocomellus sp JQ003627 Phylloporus pumilus

0.05 Figure 28. Molecular phylogenetic analysis of Xerocomellus rimosus based on ITS sequences. The evolutionary history was inferred by the Maximum Likelihood method using Jukes-Cantor model. The analysis involved 36 nucleotide sequences. There were a total of 947 positions in the final dataset. Sequences generated from LAH35036 are marked with .

69

65 AF514809_Xerocomus_chrysenteron 42 KF112357_Xerocomellus_chrysenteron AF514807_Xerocomus_chrysenteron 59 AF402139_Xerocomus_chrysenteron AF347103_Xerocomus_chrysenteron 90 AF514808_Xerocomus_chrysenteron AF402140_Xerocomus_pruinatus 35 Xerocomellus rimosus SJ65 KF030277_Xerocomellus_cf._porosp

51 KF030276_Xerocomellus_chrysenter 23 93 KF030275_Xerocomellus_zelleri DQ534625_Xerocomus_truncatus 62 66 AF050645_Xerocomus_porosporus 96 EF173308_Octaviania_columellifer 66 EF183535_Octaviania_columellifer EF183544_Octaviania_columellifer 64 EF183533_Octaviania_columellifer 96 EF173306_Octaviania_columellifer 31 EF183541_Octaviania_columellifer EF183543_Octaviania_columellifer 64 EF183539_Octaviania_columellifer EF183537_Octaviania_columellifer 100 AF514820_Xerocomus_fennicus AF514821_Xerocomus_fennicus 93 AF514818_Xerocomus_ripariellus AF050649_Xerocomus_rubellus 81 65 AF514816_Xerocomus_ripariellus 82 AF514827_Xerocomus_pruinatus 87 AF514825_Xerocomus_pruinatus 62 AF050644_Xerocomus_pruinatus KF030272_Xerocomellus_cf._chrysenteron 99 KF030273_Xerocomus_truncatus 78 AF071537_Xerocomus_chrysenteron 25 AY612842_Xerocomus_spadiceus AF514815_Xerocomus_cisalpinus 64 KF030274_Xerocomellus_cisalpinus AF514813_Xerocomus_cisalpinus 90 AF514812_Xerocomus_cisalpinus JQ924322_Xerocomellus_cisalpinus HQ161873_Tylopilus_balloui

0.01 Figure 29. Molecular phylogenetic analysis of Xerocomellus rimosus based on LSU sequences. The evolutionary history was inferred by the Maximum Likelihood method using General Time Reversible model. The analysis involved 40 nucleotide sequences. There were a total of 875 positions in the final dataset. Sequences generated from LAH35036 are marked with .

70

Genus Cortinarius

Cortinarius corrosus Fr., Epicr. syst. mycol. (Upsaliae): 266 (1838) (Figures 30 & 31)

Pileus up to 3.2–5 cm broad, convex to flat at maturity; cuticle orangish brown (7.5YR5/8) to cream (2.5Y8/6) or off white (5Y9/2), smooth and shiny; margins smooth, incurved and rimose at maturity. Lamellae light brown (5YR7/8) to grayish (10R4/4), up to 0.35 cm broad, adnexed, sub distant to close, edges entire. Lamellulae two tiered 1/2 and 1/3 of the lemellae. Cortina in the form of thin threads. Stipe up to 3.5 × 1.5 cm, apex slightly narrow and base with a marginate bulb up to 1.8 cm in diameter, central, straight to bent, surface smooth to fibrillose, orangish yellow (5YR7/10) towards the base, becoming cream (5Y9/4) towards the apex, context off white to cream, cortina remnants filamentous, brownish (10R4/12) forming a ring on the stipe.

Basidiospores [30/2/2] (11.6) 12.04–15.34 (16.04) × (6.1) 6.4–8.5 (9.44) µm, Q = (0.3) 0.34–0.47 (0.51), avQ = 0.42 ± 0.05; amygdaliform, exine verrucose, apiculus prominent, brownish,in 5% KOH. Basidia (28.3) 29.3–36.1 (37.7) × (9.4) 9.9–10.6 (11.3) µm, clavate having 2–4 starigmata, thin walled up to 3 µm, densely guttulate, hyaline 5% KOH. Marginal cells 19.8–24.5 × 7.7–9.4 µm, cylindrical to fusiform, hyaline in 5% KOH. Partial veil remnants hyphae (2.1) 2.6–3.1 (3.5) µm wide, thin, filamentous, infrequently septate, clamp connections common. Pileipellis (5.0) 4.0–5.4 (8.0) µm in diameter, filamentous, branched, septate, clamp connections frequent, pale yellow in 5% KOH. Stipitipellis hyphae (3.3) 3.8–4.2 (4.7) µm wide, filamentous, branched, hyaline in 5% KOH, septate, clamp connections frequent.

Material examined: Pakistan, Khyber Pakhtunkhwa province, Malakand division, Swat district, Kalam, 2500 m asl, on soil under Cedrus deodara, 3 Sep 2013, Sana Jabeen & Abdul Nasir Khalid K2-8; SJ41 (LAH35035); Swat district; Mashkun, 2500 m asl, on soil under C. deodara, 5 Sep 2013, Sana Jabeen & Abdul Nasir Khalid MT2OR (LAH35036).

Molecular phylogenetic characterization (Figure 32)

PCR products from ITS region of Cortinarius corrosus yielded 707 base pairs by using ITS1F and ITS4 primers. A consensus sequence of 590 base pairs was obtained by trimming the conserved motifs on both ends and BLAST searched at NCBI. It showed 99% identity to C. corrosus sequences (FJ039641, FJ039642 & DQ66381) from Canada

71 and Denmark with 100% query cover. It also showed 99% sequence identity to C. calojanthenus (KF732538) from Finland with 100% query cover and 0.0 E value.

Sequences from the closely related taxa were retrieved from the GenBank to reconstruct phylogeny. Cortinarius glaucopus (Schaeff.) Fr. (AY174787) was chosen to root the phylogenetic tree. Cortinarius corrosus clustered in a clade with other C. corrosus from Europe forming a sister clade with C. largentii Ammirati & M. M. Moser with 80% boot strap value.

Comments

The genus Cortinarius (Pers.) Gray is suspected to be the largest genus of fungi containing more than 2000 species, distributed worldwide (Kirk et al., 2008; Harrower et al., 2015; Borovička et al., 2015). Members of the genus are found in association with trees (Kuo, 2011). From Pakistan, four species have been reported till date (Ahmad et al., 1997). Cortinarius corrosus is mostly reported from Europe (Breitenbach & Kränzlin, 2000). Records also have been found from Australia (May, 2003), USA (Overholts, 1919), West Asia (Sesli, 2007). It has been found first time from South Asia during this study.

72

A

B

Figure 30. Morphology of Cortinarius corrosus. A & B. Basidiomata. A. LAH35035; B. LAH35036. Bars: A & B = 1cm.

73

Figure 31. Anatomy of Cortinarius corrosus. A–E. LAH35035. A. Basidia, basidiole and marginal cell; B. Basidiospores; C. Pileipellis; D. Stipitipellis; E. Hyphae from cortina. Bars: A = 7 µm; B = 3.3 µm; C = 10.5 µm; D = 7.8 µm; E = 0.8 µm.

74

KR674106_Cortinarius_albidolilac KR674105_Cortinarius_albidolilac 99 KR674104_Cortinarius_albidolilac KR674102_Cortinarius_albidolilac 71 KR674103_Cortinarius_albidolilac EU057055_Cortinarius_lilacinovel EU655668_Cortinarius_lilacinovel 77 99 EU056960_Cortinarius_lilacinovel 70 95 EU056959_Cortinarius_lilacinovel 99 KF673472_Cortinarius_cobaltinus KF673471_Cortinarius_cobaltinus 17 96 EU056969_Cortinarius_spectabilis 93 DQ663241_Cortinarius_caesiocinct 98 EU056978_Cortinarius_calochrous 95 DQ663234_Cortinarius_calochrous EU056977_Cortinarius_calochrous KF732347_Cortinarius_metarius 89 42 EU057053_Cortinarius_barbarorum EU655671_Cortinarius_barbarorum 99 DQ663236_Cortinarius_barbarorum DQ663238_Cortinarius_barbarorum Calochori EU057059_Cortinarius_largentii DQ323964_Cortinarius_corrosus EU056974_Cortinarius_corrosus 80 Cortinarius_corrosus SJ41 FJ039642_Cortinarius_corrosus KF732272_Cortinarius_calojanthin 76 FJ039641_Cortinarius_corrosus DQ663281_Cortinarius_corrosus KF732538_Cortinarius_calojanthin EU056975_Cortinarius_corrosus EU057057_Cortinarius_corrosus KF732561_Cortinarius_sublilacino 99 EU655674_Cortinarius_sublilacino EU056988_Cortinarius_sublilacino EU056987_Cortinarius_albovestitu 59 62 EU057056_Cortinarius_haasii EU655684_Cortinarius_aurantioruf 99 AY669561_Cortinarius_haasii EU056985_Cortinarius_haasii 67 EU056986_Cortinarius_haasii AY174787_Cortinarius_glaucopus Out group

0.01 Figure 32. Molecular phylogenetic analysis of Cortinarius corrosus based on ITS sequences. The evolutionary history was inferred by the Maximum Likelihood method using Jukes-Cantor model. The analysis involved 42 nucleotide sequences. There were a total of 639 positions in the final dataset. Sequences generated from LAH35035 are marked with .

75

Cortinarius longistipus nom. prov. (Figures 33 & 34)

Etymology: The specific epithet refers to the long stipe of the taxon under study.

Pileus up to 4.5 cm broad, convex to flat at maturity; cuticle orangish (10R5/14) to dark brown (10 R5/14) having blacking margins (5YR1/2) becoming whole surface blakish brown (5YR1/2) when mature, surface dry, smooth and shiny; margins smooth, incurved and rimose at maturity. Lamellae dark brown (10R3/8), up to 0.5 cm broad, adnexed, sub distant to distant, edges entire. Lamellulae absent. Cortina webbed. Stipe up to 13 × 1.6 cm, apex slightly wide up to 1.9 cm in diameter, base narrow up to 0.6 cm in diameter, central, straight to bent, surface smooth to fibrillose, orangish (10R5/12) towards the base, becoming cream (5Y9/4) towards the apex, context off white to cream.

Basidiospores [30/2/2] (6.1) 9.2–9.9 (10.8) × (5.4) 5.7–6.3 (6.4) µm, Q = (0.10) 0.13–0.15 (0.16), avQ = 0.14; subglobose to elliptical, exine verrucose, apiculus prominent, brownish in 5% KOH. Basidia (28.7) 33.0–34.6 (39.4) × (8.4) 8.9–10 (10.6) µm, clavate having 2–4 starigmata, thin walled, wall up to 3 µm thick, hyaline 5% KOH. Marginal cells (16.2) 22.4–33.9 (34.9) × (5.2) 7.5–8.3 (8.7) µm, long cylindrical to fusiform, hyaline in 5% KOH. Partial veil remnants hyphae (2.83) 3.0–3.5 (4.2) µm wide, branched, septate, clamped septa frequent, purplish in 5% KOH. Pileipellis hyphae 3–5.6 µm in diameter, infrequently branched, septate. Stipitipellis hyphae (4.0) 4.7–8.2 (11) µm wide, filamentous, branched, hyaline in 5% KOH, septate, clamped septa infrequent.

Material examined: Pakistan, Khyber Pakhtunkhwa province, Hazara division, Mansehra district, Khanian, 2500 m asl, on soil under Cedrus deodara, 5 Aug 2014, Sana Jabeen & Abdul Nasir Khalid KH3; SJ101 (LAH35033); Malakand division, Swat district, Mashkun, 2500 m asl, on soil under C. deodara, 5 Sep 2013, Sana Jabeen & Abdul Nasir Khalid K4-25; SJ10 (LAH35034).

Molecular phylogenetic characterization (Figure 35)

Sequencing of the PCR products of ITS region of Cortinarius longistipus yielded 615–714 base pairs by using ITS1F and ITS4 primers. Consensus sequence of 532 base pairs was obtained by trimming the motif and BLAST searched at NCBI. It showed 99% identity to C. bulliardi (Pers.) Fr. sequence from Germany and C. cinnabarinus Fr. sequences (JX114963 & AY669662) from Finland and Germany with 100% query cover and 0.0 E value.

76

To reconstruct phylogeny, closely related sequences of the ITS region were retrieved from the GenBank. Taxa from section Calochori were chosen as out groups. Cortinarius longistipus clustered in section Cinnabarini within a clade including C. bulliardii and C. cinnabarinus with 88% boot strap value. Cortinarius longistipus separated from C. bulliardii and C. cinnabarinus forming a sister lineage with C. cinnabarinus.

Comments

Cortinarius longistipus nom. prov. is characterized by its dark brown to blackish pileus, dark brown lamellae and a long stipe which is narrow at the base. The species is a close relative of C. bulliardii and C. cinnabarinus. It is distinct morphologically from these taxa by its pileal color, size and shape of the stipe. Cortinarius bulliardi and C. cinnabarinus bear a comparatively short stipe with a bulbous base. Moreover the pileus color of C. bulliardi is red or reddish pink and it is cinnabar to reddish in C. cinnabarinus. C. cinnabarinus also differs from C. longistipus by the presence of a low umbo (Fries, 1838; Ammirati et al., 2013). Molecular pylogenetic analysis based on ITS sequences also supports C. longistipus as a distinct species with a strong boot strap value.

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A

C

B D Figure 33. Morphology of Cortinarius longistipus. A–D. Basidiomata. A & B. LAH35034; C & D. LAH35033 (holotype). Bars: A & B = 1 cm; C & D = 2.5 cm.

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Figure 34. Anatomy of Cortinarius longistipus. A–G. LAH35033. A. Basidiospores; B. Basidia C & D. Basidioles with marginal cells; E. Hyphae from cortina; F. Pileipellis; G. Stipitipellis. Bars: A = 5.5 µm; B = 10 µm; C & D = 8 µm; E–G = 10 µm.

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KJ206490_Cortinarius_nolaneiform KJ206486_Cortinarius_nolaneiform 100 KJ206491_Cortinarius_nolaneiform KJ206487_Cortinarius_nolaneiform KJ206489_Cortinarius_nolaneiform 45 KJ206488_Cortinarius_nolaneiform KJ206509_Cortinarius_uraceomajal KJ206514_Cortinarius_uraceomajal 72 KJ206513_Cortinarius_uraceomajal 99 KJ206512_Cortinarius_uraceomajal KJ206484_Cortinarius_colymbadinu KJ206482_Cortinarius_colymbadinu KJ206483_Cortinarius_colymbadinu 43 99 JF907865_Cortinarius_colymbadinu KC842404_Cortinarius_colymbadinu JF907860_Cortinarius_bulliardii 100 AF389154_Cortinarius_bulliardii JX114942_Cortinarius_bulliardii 90 Cortinarius longistipus SJ101b Cinnabarini 100 37 Cortinarius longistipus SJ101 88 Cortonarius longistipus SJ10 AY669662_Cortinarius_cinnabarinu 59 JX114943_Cortinarius_cinnabarinu JX114944_Cortinarius_cinnabarinu 81 KC842405_Cortinarius_cinnabarinu GQ159845_Cortinarius_californicu 67 GQ159788_Cortinarius_californicu 99 FJ039679_Cortinarius_californicu 12 GQ159895_Cortinarius_californicu KJ206518_Cortinarius_uraceonemor 100 KJ206519_Cortinarius_uraceonemor KJ206521_Cortinarius_uraceonemor KJ206515_Cortinarius_uraceonemor KC608598_Cortinarius_uraceus 87 KC608595_Cortinarius_uraceus JX407339_Cortinarius_uraceus 91 KJ206524_Cortinarius_uraceus DQ120758_Cortinarius_anisatus 34 DQ120757_Cortinarius_anisatus 100 DQ120756_Cortinarius_anisatus DQ120754_Cortinarius_anisatus DQ120755_Cortinarius_anisatus JX407305_Cortinarius_anisochrous 100 JX407304_Cortinarius_anisochrous 92 JX407307_Cortinarius_anisochrous 81 JX407316_Cortinarius_fuscobovina JX407318_Cortinarius_fuscobovina 100 KM586144_Cortinarius_fuscobovinu 44 JX407321_Cortinarius_fuscobovinu JX407322_Cortinarius_fuscobovinu JX407320_Cortinarius_fuscobovinu 100 JX407266_Cortinarius_bovinaster Bovini 60 JX407265_Cortinarius_bovinaster 80 Cortinarius oulankaensis SA70 99 FJ039672_Cortinarius_oulankaensi 74 JX407296_Cortinarius_oulankaensi JX407295_Cortinarius_oulankaensi 100 100 KC905157_Cortinarius_bovarius 43 KC905156_Cortinarius_bovarius KC905158_Cortinarius_bovarius JX407285_Cortinarius_bovinus 71 JX407288_Cortinarius_bovinus JX407271_Cortinarius_bovinus 98 JX407287_Cortinarius_bovinus JX407283_Cortinarius_bovinus 82 JX407289_Cortinarius_bovinus JX407284_Cortinarius_bovinus DQ663250_Cortinarius_calochrous DQ663237_Cortinarius barbarorum Out group 100

0.02 Figure 35. Molecular phylogenetic analysis of Cortinarius longistipus based on ITS sequences. The evolutionary history was inferred by the Maximum Likelihood method using Jukes-Cantor model. The analysis involved 69 nucleotide sequences. There were a total of 658 positions in the final dataset. Sequences generated from fruit bodies of Cortinarius longistipus are marked with .

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Genus Geastrum

Geastrum galiyensis nom. prov. (Figures 36 & 37) Mature basidiomata globose to depressed globose, sessile, up to 4.5 cm in diameter, when mature, splitting in a stellate pattern, encrusted with soil particles and other debris material. Peridium double. Exoperidium up to 1.5 mm thick, splitting in to 8 acute rays; rays expanded when fresh, recurved inward upon drying; composed of three layers; outer layer thin grayish brown (2.5YR2/4) to off white, non persistent; middle layer light brown (7.5YR6/6); light brown to dark brown (7.5YR3/6) with age, thick up to 2 mm, extensively cracked in mature specimens. Endoperidium sessile, grayish brown (10YR4/4), opens by a pore; pore prominent, area around the pore is off white, upto 4 mm in radius. brownish, cottony. Basidiospores [20/2/1] (3.0) 3.4–4.6 (4.9) × (2.9) 3.9–4.5 (4.7) µm, Q = (0.83) 0.85–0.97 (1.0), avQ = 0.92, globose to sub globose, echinulate, , apiculus prominent, brownish in 5% KOH. Hyphae from gleba (2.1) 3.9–4.5 (5.3) µm, infrequently septate, brownish in 5% KOH.

Etymology: The specific epithet refers to the site of collection "Kuza Gali".

Material examined: Pakistan, Punjab province, Rawalpindi division & district, Kuza Gali, 2500 m asl, on soil under Cedrus deodara, 21 Sep 2013, Sana Jabeen & Hassan Ayub KG-4; SJ61 (LAH35036).

Molecular phylogenetic characterization (Figure 38)

Sequencing of the PCR products of ITS region of Geastrum galiyensis yielded 606 & 607 base pairs by using ITS1F and ITS4 primers. Consensus sequence of 517 base pairs was obtained by trimming at the conserve sites. BLAST searched revealed that the sequence showed 84% identity to G. saccatum Fr. sequences (KC581968, KC581969 & KC581967) from Sweden with 94% query cover.

To reconstruct phylogeny, sequences of the ITS region of closely related taxa were retrieved from the GenBank. Myriostoma coliforme (Dicks.) Corda (KF988337) was chosen as out group. The sequence generated during this study clustered in a clade including , G. aff. lageniforme and an indentified Geastrum species. It got separated from all these taxa forming a lineage of its own.

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Comments

Geastrum galiyensis nom. prov. is morphologically characterized by its stellate fruit body having 8 rays, grayish brown non persistent outer layer and dark brown inner layer of exoparidium, a prominent pore having an off white disc around the pore. It is comparable with G. saccatum. Outer surface of G. saccatum is pinkish tan to yellowish brown and inner layer is off white to pinkish (Sundberg & Bessette, 1987). In molecular analysis based on ITS sequences revealed that the taxon differs from other reported taxa. It gets separated along with another undescribed taxon of Geastrum (KF988452 & KF988453) from Argentina forming a sister clade with G. saccatum and G. aff lageniforme from Sweden and Argentina respectively.

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Figure 36. Morphology of Geastrum galiyensis. Basidiomata LAH35036 (holotype). Bar = 1.5 cm

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Figure 37. Anatomy of Geastrum galiyensis. A & B. LAH35036. A. Basidiospores; B. Hyphae from gleba. Bars: A = 4.5 µm; B = 6.8 µm.

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JN845112 Geastrum pectinatum JN845111 Geastrum pectinatum 24 KC581962 Geastrum pectinatum KP687517 Geastrum pectinatum KP687516 Geastrum pectinatum 26 KF988413 Geastrum pectinatum EU784239 Geastrum pectinatum EU784241 Geastrum pectinatum NR 132896 Geastrum meridionale KP687512 Geastrum meridionale 45 KF988412 Geastrum meridionale KP687508 Geastrum meridionale 97 KF988433 Geastrum saccatum KF988432 Geastrum saccatum 18 KC581975 Geastrum xerophilum 38 16 KF988383 Geastrum aff hariotii KF988462 Geastrum sp KF988419 Geastrum pseudolimbatum 14 KF988420 Geastrum pseudolimbatum 93 KC581974 Geastrum pseudolimbatum KC581973 Geastrum pseudolimbatum 91 NR 132895 Geastrum thanatophilum KF988364 Geastrum thanatophilum KF988434 Geastrum schmidelii 40 KC582006 Geastrum schmidelii JN845122 Geastrum schmidelii 91 JN845121 Geastrum schmidelii EU784247 Geastrum schmidelii KF988458 Geastrum sp KF988459 Geastrum sp 42 KJ127024 Geastrum sp KJ127023 Geastrum sp KJ127025 Geastrum sp KF988395 Trichaster melanocephal KF988377 Geastrum fuscogleba 34 KF988396 Trichaster melanocephal 10 KC581982 Trichaster melanocephal 99 KF988452 Geastrum sp 16 KF988453 Geastrum sp Geastrum galiyensis SJ61 91 KF988392 Geastrum aff lageniform KF988391 Geastrum aff lageniform 93 KF988394 Geastrum aff lageniform 26 KF988393 Geastrum aff lageniform 40 KC581967 Geastrum saccatum KC581968 Geastrum saccatum 75 49 KC581969 Geastrum saccatum KF988339 Geastrum lageniforme KF988388 Geastrum lageniforme 93 KC581966 Geastrum lageniforme KF988337 Myriostoma coliforme Out group

0.02 Figure 38. Molecular phylogenetic analysis of Geastrum galiyensis based on ITS sequences. The evolutionary history was inferred by the Maximum Likelihood method using Jukes-Cantor model. The analysis involved 51 nucleotide sequences. There were a total of 377 positions in the final dataset. Sequences generated from LAH35036 are marked with .

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Genus Gomphidius

Gomphidius flavostipus nom. prov. (Figures 39 & 40)

Etymology: The specific epithet refers to the yellow color of stipe.

Pileus up to 5 cm broad, convex to plano convex at maturity; cuticle off white (5Y9/2) to brownish (2.5YR5/10) or grayish (5YR4/2) becoming brownish black at maturity (5YR2/6) in the form of patches on off white context; surface dry, smooth and shiny, sometimes cracked and the off white context visible in between the cracks; margins dentate and incurved. Lamellae light gray (2.5Y8/2) to dark gray (2.5Y4/2), broad up to 0.5 cm, decurrent, distant, edges wavy. Lamellulae versatile in length. Stipe up to 4.5 × 1.2 cm, apex slightly wide up to 1.5 cm in diameter becoming narrower towards the base, base up to 0.4 cm in diameter, central, straight to bent, off white to grayish brown (5YR5/2) from the top becoming bright yellow (5Y8/10) towards the base.

Basidiospores [30/2/2] (14.7) 16.4–16.9 (17.2) × (6.1) 6.5–7.2 (7.3) µm, Q = 2.2– 2.7, avQ = 2.5; narrowly ellipsoid to fusoid, smooth, apiculus prominent, dark brown in 5% KOH. Basidia 43.25–53.98 × 8.51–10.34 µm, clavate having 2–4 starigmata, thin walled, wall up to 2.8 µm thick, hyaline 5% KOH. Cystidia 40.30–60.2 × 8.3–10.23 µm, long cylindrical to fusiform, hyaline in 5% KOH. Pileipellis hyphae 2.58–9.12 µm in diameter, infrequently branched, septate, clavate to fusoid terminals. Stipitipellis hyphae 3.68–9.37 µm wide, filamentous, branched, septate, hyaline in 5% KOH.

Material examined: Pakistan, Khyber Pakhtunkhwa province, Malakand division, Swat district, Kalam 2400 m asl, on soil under Cedrus deodara, 3 Sep 2013, Abdul Nasir Khalid K2-9; (LAH35038); K2-18; SJ11 (LAH35039); Mashkun 2500 m asl, on soil under C. deodara, 5 Sep 2013, Sana Jabeen K4-30; SJ68 (LAH35040).

Molecular phylogenetic characterization (Figures 41 & 42)

Sequencing of the PCR products of ITS region of Gomphidius flavostipus yielded 744–768 base pairs by using ITS1F and ITS4 primers. Consensus sequence of 660 base pairs was obtained by trimming at conserved motifs and BLAST searched at NCBI. It showed 93% identity to G. oregonensis Peck sequences (KC581306, DQ533976 & L54114) from Canada and USA. It also showed 92–93% identity to G. glutinosus (Schaeff.) Fr. from Czech Republic and USA with 99–100% identity and 0.0 E value.

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Sequencing of the PCR products of LSU yielded 697–879 base pairs by using LR0R and LR5 primers. Consensus sequence of 879 base pairs was BLAST searched at NCBI. It was found 94% identical to G. oregonensis (AF071530) from USA with 89% query cover and 94% identical to G. glutinosus (AF098398) from Germany with 87% query cover and 0.0 E value.

To reconstruct phylogeny, closely related sequences of the ITS and LSU regions were retrieved from the GenBank. Suillus spp. (JQ958317 & JQ958319) were chosen as out group taxa in ITS data set while Russula acrifolia Romagn. (DQ4211998) was chosen in LSU data set. Gomphidius flavostipus clustered with other taxa of the same genus in Gomphidius clade. It gets separated from all other taxa forming a sister clade with 83% and 58% boot strap value in ITS and LSU data analysis respectively.

Comments

Gomphidius flavostipus nom. prov. is characterized by its off white to grayish brown or blackish pileal surface in the form of patches, grayish decurrent lamellae, bright yellow narrow stipe base and narrowly ellipsoid dark brown basidiospores. It can be compared with G. glutinosus and G. oregonensis. Gomphidus glutinous differs from G. flavostipus by its purplish sticky gelatinous pileus and spindle shaped basidiospores (Zeitlmayr, 1976; Phillips, 2006). Gomphidius oregonensis, a closely related taxon, is more pinkish and sticky at its earlier stage of development (Kuo, 2014). Its spores are brownish in KOH and are smaller (<10 µm), while G. flavostipus is dry with slightly elastic contest and bears dark brown and comparatively larger basidiospores. Molecular pylogenetic analysis based on ITS sequences also supports that G. flavostipus is distinct from other reported taxa. Only a few sequences of LSU from Gomphidius spp. are available in GenBank. It also formed its own lineage within LSU sequences of the genus.

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A

B Figure 39. Morphology of Gomphidius flavostipus. A & B. Basidiomata. A. LAH35040 (holotype); B. LAH35039. Bars: A = 0.8 cm; B = 0.7 cm.

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Figure 40. Anatomy of Gomphidius flavostipus. A–E. LAH35040. A. Basidiospores; B. Cystidia; C. Basidia and basidiole; D. Pileipellis; E. Stipitipellis. Bars: A–E = 10 µm.

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DQ53397445_Chroogomphus_vinicolo 89 AF20564520_Chroogomphus_vinicolo 60 L540951_Chroogomphus_vinicolor 59 AF2056466_Chroogomphus_vinicolor 100 AF2056601_Chroogomphus_jamaicens 48 AF2056581_Chroogomphus_jamaicens FJ51332649_Chroogomphus_filiform 32 EU70632350_Chroogomphus_filiform 99 51 FJ51332749_Chroogomphus_filiform KC1520792_Chroogomphus_jamaicens 26 DQ36789460_Chroogomphus_rutilus FJ157000_Chroogomphus_leptocysti Chroogomphus 99 EU70632945_Chroogomphus_roseolus 97 EF42362012_Chroogomphus_roseolus

58 66 AF2056504_Chroogomphus_helveticu AF20564215_Chroogomphus_helvetic 87 GU18751447_Chroogomphus_helvetic 100 EF4236235_Chroogomphus_confusus 90 EF42362213_Chroogomphus_confusus 98 EU69724010_Brauniellula_albipes 81 EU69723919_Brauniellula_albipes 88 AF2056375_Brauniellula_albipes AF20564818_Chroogomphus_tomentosus 100 KC5813211_Chroogomphus_tomentosus 100 AF2056591_Gomphidius_nigricans AY0774744_Gomphidius_nigricans 96 Gomphidius flavostipus SJ11b 95 Gomphidius flavostipus SJ11 93 Gomphidius flavostipus SJ68 100 AY0774721_Gomphidius_glutinosus 44 AF20564721_Gomphidius_glutinosus 83 EU79157848_Gomphidius_sp Gomphidius KC5813061_Gomphidius_oregonensis 99 49 L541141_Gomphidius_oregonensis DQ53397645_Gomphidius_oregonensi 81 EF5309419_Gomphidius_smithii 16 AY07747125_Gomphidius_smithii 80 DQ099900_Gomphidius_subroseus 100 EU5970907_Gomphidius_cf_subroseu 84 DQ384576_Gomphidius_subroseus JQ958319_Suillus_ponderosus Out group 100 JQ958317_Suillus_caerulescens

0.02 Figure 41. Molecular phylogenetic analysis of Gomphidius flavostipus based on ITS sequences. The evolutionary history was inferred by the Maximum Likelihood method using Jukes-Cantor model. The analysis involved 42 nucleotide sequences. There were a total of 734 positions in the final dataset. Sequences generated from from G. flavostipus are marked with .

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64 AB284601_Suillus_pictus 87 AY684154_Suillus_pictus 60 AY612830_Suillus_pictus AY612829_Suillus_americanus 51 83 AB284491_Suillus_americanus AB284482_Suillus_sp 43 AY612825_Suillus_luteus 100 Suillus AF042622_Suillus_luteus AB284490_Suillus_placidus 100 AB284485_Suillus_placidus_ KF112430_Suillus_aff._luteus 70 65 GU187598_Suillus_bresadolae 98 AB284478_Suillus_laricinus KJ146731_Suillus_lakei KC581306_Gomphidius_oregonensis AF071530_Gomphidius_oregonensis

27 58 Gomphidius flavostipus SJ11 Gomphidius AF098398_Gomphidius_glutinosus 37 EF530941_Gomphidius_smithii 91 DQ384576_Gomphidius_subroseus

99 KC581321_Chroogomphus_tomentosus 53 HM240518_Chroogomphus_tomentosus DQ534669_Gomphidius_roseus

34 AF071529_Chroogomphus_vinicolor Chrogomphus AY586645_Chroogomphus_rutilus 65 EU852806_Brauniellula_albipes 81 AF098399_Chroogomphus_rutilus DQ421998_Russula_acrifolia Out group

0.02 Figure 42. Molecular phylogenetic analysis of Gomphidius flavostipus based on LSU sequences. The evolutionary history was inferred by the Maximum Likelihood method using Jukes-Cantor model. The analysis involved 28 nucleotide sequences. There were a total of 445 positions in the final dataset. Sequences generated from from G. flavostipus are marked with .

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Genus Hebeloma

Hebeloma angustisporium Hesler, Kew Bull. 31: 471 (1977) (Figures 43 & 44)

Pileus 1.2–4 cm broad, convex when young, becoming flat at maturity, surface dry and cracked, context off white (5Y9/2) from within the cracks, cuticle light brown (7.5YR7/6 to 10YR7/6); margins smooth, incurved, rimose at maturity. Lamellae up to 0. 3 cm broad, light brown (10YR7/6), adnexed, sub distant to close, edges slightly dentate. Lamellulae versatile lengths, 4 tiered, alternating with lamellae. Stipe 2.5–3.5 cm × 0.3–1 cm, central, cylindrical to bent, slightly narrower towards the base; surface granular to fibrillose, off white (5Y9/2) with some brownish (7.5YR7/6) patches.

Basidiospores [30/2/1] (8.4) 8.6–10.57 (11.1) × (5.0) 5.2–6.0 (6.2) µm, Q = (1.4) 1.6–1.8 (2), avQ = 1.8; amygdaliform, ornamented with low verrucae , apiculus prominent, brown in 5% KOH. Basidia 22.4–32.3 × 8.0–8.8 µm, clavate having 2–4 starigmata, thin walled, densely guttulate, hyaline 5% KOH. Cystidia 18.7–23.4 × 5.8–7.2 µm, clavate, hyaline in 5% KOH. Pileipellis hyphae (1.8) 2.3–4.1 (4.5) µm in diameter, filamentous, infrequently branched, septate, clavate to fusoid terminals, hyaline in 5% KOH. Stipitipellis hyphae (1.3) 2.2–2.5 (3.8) µm wide, filamentous, infrequently branched, septate, clavate to fusoid terminals, hyaline in 5% KOH.

Material examined: Pakistan, Khyber Pakhtunkhwa province, Malakand division, Swat district, Kalam 2400 m asl, on soil under Cedrus deodara, 3 Sep 2013, Sana Jabeen & Abdul Nasir Khalid K2-10; SJ40 (LAH35041).

Molecular phylogenetic characterization (Figure 45)

Sequences of 709–712 base pairs were obtained from the PCR products of ITS region of Hebeloma angustisporium using ITS1F and ITS4 primers. A consensus sequence of 660 base pairs was created and trimmed at conserved motifs. It was BLAST searched at NCBI. It showed 99% identity to H. naviculosporum Heykoop, G. Moreno & Esteve-Rav. from Germany (KT071039), Slovakia (KT071041) and Spain (KT071041) with 100% query cover. It also showed 99% identity to H. truncatum (Schaeff.) P. Kumm (FJ168588) from France with 100% query cove and 98% identity. It was also found 98% identical to H. angustisporium (HQ179476) and H. pallidum P. Kumm. (JF908039) from USA and Italy with 98 and 100% query cover respectively.

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To reconstruct phylogeny, closely related ITS sequences were retrieved from the GenBank. pseudocamerina Singer (AJ585508), Gymnopilus penetrans (Fr.) Murrill (AF325663) and Hebelomina neerlandica Huijsman (AY320399) were chosen as out group (Boyle et al., 2006). Sequences of Hebeloma angustisporium clustered with H. angustisporium sequence of type specimen from USA with strong boot strap value in section Sacchariolentia (J. E. Lange ex M. Bon) H. Boyle.

Comments

Hebeloma angustisporium was first collected by Hesler in 1959 from Tennessee, USA and described in 1977 (Hesler, 1977). The species is a close relative of H. sacchariolens Quél and H. tomentosum (M.M. Moser) Gröger & Zschiesch, forming a sister clade with these taxa in section Sacchariolentia, a new combination of subsection Sacchariolentia J. E. Lange ex M. Bon, based on the distinct phylogenetic position of this group (Boyle et al., 2006). In present study H. angustisporium, clustred with the type specimen sequence along with H. pallidum from Itlay. Other H. pallidum sequnces along with its synonyms are clustered in /Indusiata clade (Vesterholt, 1989). So, H. pallidum from Italy clustered with H. angustisporium sequences could be the H. angustisporium. Occurrence of H. angustisporium in Pakistan is an addition to the fungi of Pakistan.

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Figure 43. Morphology of Hebeloma angustisporium. Basidiomata LAH35041. Bar = 1.5 cm.

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Figure 44. Anatomy of Hebeloma angustisporium. A–D. LAH35041. A. Basidiospores; B. Basidia and cystidia; C. Pileipellis; D. Stipitipellis. Bars: A–D = 10 µm.

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AY311519 H malenconii 59 AY312976 H pallidum AY320397 H versipelle AY311521 H mesophaeum var crassipes AY309962 H collariatum 39 AY320395 H testaceum AY312980 H psammophilum AF124692 H mesophaeum / Indusiata /Hebeloma AY312984 H repandum AY308585 H psammophilum 63 AY311520 H marginatulum AY311520 H marginatulum AY30995897 H bruchetii 89 AY309958 H bruchetii 84 96 DQ007992 H cistophilum DQ007993 H cistophilum /Indusiata /Cistophilum AY312983 H remyi GU234160 H sp 87 AY311523 H monticola 53 AY312977 H polare AY311522 H aff nigellum 47 AY308586 H atrobrunneum /Indusiata /Amygdalina AY312986 H saliciphilum AY308586 H atrobrunneum 75 AY311524 H nigellum GU234097 H kuehneri KJ146711 H leucosarx 40 AB211268 H leucosarx AY308582 H aestivale KM576603 H sp KR01980784 H leucosarx AY320392 H stenocystis AY818351 H velutipes KC110680 H velutipes EU570175 H velutipes EU379675 H crustulinifo /Velutipes 73 KC110675 H velutipes JQ751200 H velutipes KC110678 H velutipes KF639965 H incarnatulum KC110672 H velutipes AF124684 H incarnatulum AF430291 H incarnatulum EU570173 H velutipes AF124682 H sinapizans AF124701 H truncatum AY320380 H sinapizans /Sinapizantia 99 AY320380 H sinapizans KC603716 H radicosum KC477649 H radicosum /Myxocybe 100 FJ168582 H radicosum 91 AY311526 H pallidoluctuosum 80 AF124680 H tomentosum AF124689 H sacchariolens AY312985 H sacchariolens A 99 JF908039 H pallidum B /Sacchariolentia HQ179476 H angustisporium H angustisporium SJ40 84 H angustisporium SA40 98 AF124715 H sarcophyllum AB742193 H porphyrospora AB742175 H vinosophyllum 46 AB742177 H vinosophyllum /Porphyrospora AB742184 H vinosophyllum 98 AB742185 H vinosophyllum GU591648 H aminophilum 82 GU591646 H aminophilum GU591649 H aminophilum /Porphyrospora /Aminophilum 47 GU591644 H aminophilum f hygrosa GU591643 H aminophilum f hygrosa GU591640 H youngii 59 AY818352 Anamika lactariolens 36 AF407163 Anamika indica /Porphyrospora /Anamika 34 AY575917 Anamika angustilamellat 89 GU591634 H subvictoriense GU591635 H khogianum 99 GU591636 H victoriense /Porphyrospora /Victoriense GU591637 H victoriense 96 GU591638 H victoriense 95 GU591655 H mediorufum 100 GU591656 H mediorufum 95 GU591653 H nothofagetorum /Mediorufum GU591654 H nothofagetorum 91 GU591650 H lacteocoffeatum 94 AY320396 H vaccinum AY948192 H vaccinum 53 AF124705 H helodes AF124669 H hiemale AF124690 H helodes 61 EU887517 H cavipes 95 AF124709 H helodes AY311525 H oculatum AY309964 H fragilipes AY948191 H leucosarx /Denudata 62 AY320393 H subconcolor 68 AY309959 H brunneifolium AF124702 H pusillum 66 AY312978 H populinum AY308584 H alpinum 5969 AF124683 H crustuliniforme AF124678 H leucosarx AF124716 H crustuliniforme AY308583 H albocolossum AY320394 H subsaponaceum 90 AY309961 H calyptrosporum AF124693 H birrum internal /Myxocybe /Scabrispora AY312981 H pumilum 85 AY309963 H cylindrosporum AF124675 H danicum AY312987 H senescens /Myxocybe 77 JF908043 H truncatum EU570185 H erumpens 56 EU570182 H theobrominum /Theobromina 57 AF124699 H circinans 95 AJ585508 Galerina pseudocamerina AF325663 Gymnopilus penetrans Out group 100 99 D AY320399 Hebelomina neerlandica

0.02 Figure 45. Molecular phylogenetic analysis of Hebeloma angustisporium. based on ITS sequences. The evolutionary history was inferred by the Maximum Likelihood method using Jukes-Cantor model. The analysis involved 117 nucleotide sequences. There were a total of 483 positions in the final dataset. Sequences generated from LAH35041 are marked with .

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Genus Inocybe

Inocybe alba nom. prov. (Figures 46 & 47)

Etymology: The specific epithet refers to the whitish basidiomata of the taxon.

Pileus up to 1.8 cm wide, spherical, prominent umbo, conical, surface dry, fibrillose, whitish (10Y8/4) to brownish (7.5YR6/10), margins incurved, umbo yellowish (5Y8/6) becoming brownish (7.5YR6/10) towards maturity. Lamellae adnexed, moderately close, up to 0.2 cm broad, golden brown (10YR5/6). Stipe 5.5 × 0.4 cm, central, cylindrical, narrower towards the apex and wider towards the base, surface dry, fibrillose, context off white (5Y9/2).

Basidiospores [30/2/2] (12.7) 13.2–14.4 (16.4)  (5.3) 6.1–6.9 (8.2), Q = (1.90) 1.94–2.20 (2.50), avQ = 2.0, ellipsoid, sometimes constricted from the centre, smooth, guttulate, brown in 5% KOH. Basidia (38.1) 40.3–42.2 (43.7)  (9.2) 9.5–11.8 (12.1) µm, clavate, guttulate, observed at the base, hyaline in 5% KOH. Hymenal cystidia (20.2) 25.4–27.4 (30.3)  (4.4) 4.79–5.51 (6.53) µm, clavate, connected with basal cell, clamp connection common, hyaline in 5% KOH. Caulocystida (43.1) 45.2– 50.3 (56.11)  (7.6) 9.4–10.9 (11.2) µm, clavate, clamp connections observed at the base, hyaline in 5% KOH. Stipitipellis hyphae (6.1) 8.3–11.04 (11.69) µm wide, filamentous, rarely branched, clavate terminals, septate, clamp connections observed. Pileipellis hyphae (4.6) 5.2–6.9 (7.7) µm wide, filamentous, branched, septate, clamp connections frequent, hyaline n 5% KOH.

Material examined: Pakistan, Khyber Pakhtunkhwa province, Hazara division, Mansehra district, Khanian, 2500 m asl, on soil under Cedrus deodara, 5 Aug 2014, Sana Jabeen KH-2; SJ102 (LAH35047); Malakand division, Swat district, Kalam, 2400 m asl, on soil under C. deodara, 4 Sep 2013, Sana Jabeen KI; (LAH35045); Mashkun, 2500 m asl, on soil under C. deodara, 5 Sep 2013, Aamna Ishaq K4I; SJ147 (LAH35046) .

Molecular phylogenetic characterization (Figure 50)

Sequencing of the PCR products of nrITS region of Inocybe alba yielded 747–784 base pairs by using ITS1F and ITS4 primers. A consensus sequence of 658 base pairs was subjected to BLAST at NCBI. It showed 93 and 94% identity to I. sororia Kauffman (HQ604626 & KP783443) from USA and Russia with 93% and 87%

97 query cover, respectively. It was also found 92% identical to I. dulcamaroides Kühner (FJ904126 & FJ904127) from Sweden with 94% query cover and 0.0 E value.

To reconstruct phylogeny, closely related ITS sequences were retrieved from the GenBank. Auritella foveata C.K. Pradeep & Matheny (GU062740) was chosen as out group (Larsson et al., 2009). Sequences of Inocybe alba clustered with I. breviterincarna D.E. Stuntz ex Kropp, Matheny & Hutchison, I. dulcamaroides and I sororia same clade within sub genus Rimosae sensu stricto following classification by Larsson et al. (2009). It gets separated from all these taxa forming its own lineage within the same clade.

Comments

Inocybe alba nom. prov. is characterized by its small whitish basidiomata, fibrillose stipe, smooth and elliptical basidiospores. The species is comparable with I. dulcamaroides which bears brown basidiomata and a small wide stipe pilea disc is not prominent and it becomes convex to flat at maturity (Larsson et al., 2009). Inocybe alba has comparatively long stipe with a narrow diameter, a very prominent umbo is present which lasts with time. Inocybe breviterincarnata is another closely related species with I. alba. Althouhgh there are no much distinguishable features observed anatomically, but morphologcally I. breviterincarnata differs on the basis of its pinkish lamellae and brownish pileus as well as a pinkish to brownish stipe (Kropp, 2013). While in I. alba, whitish color is prominent throughout the basdiomata and it bears golden brown lamellae. Molecular phylogenetic analysis based on ITS sequnces also confirms its position in clade E within subgenus Rimosae following Larsson et al. (2009).

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Figure 46. Morphology of Inocybe alba. Basidiomata LAH35047 (holotype). Bar = 0.5 cm

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Figure 47. Anatomy of Inocybe alba. A–F. LAH35047. A. Basidiospores; B. Basidia; C. Caulocystidia; D. Hymenal cystidia, E. Pileipellis; F. Stipitipellis. Bars: A–F = 10 µm.

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Inocybe flavellorimosa nom. prov. (Figures 48 & 49)

Etymology: The specific epithet refers to the yellowish and rimose basidiomata.

Pileus up to 4.5 cm wide, spherical, prominent umbo, conical when young, becoming wider towards maturity, surface dry, fibrillose; margins incurved, radially rimose, yellowish brown (10YR8/8) with dark brown (10R4/12) striations. Lamellae regular, adnate to adnexed, moderately close, up to 0.25 cm broad, yellowish brown (10YR7/8), becoming dark brown (10R6/14) towards maturity. Lamellulae versatile in length, alternating with lamellae. Stipe 7.5 × 0.7 cm, central, cylindrical, narrower towards the apex and wider towards the base, base rimose, surface dry, context off white under a yellowish (10YR8/8) to dark brown (5YR2/6) stipitipellis, stipitipellis peeling off from the stipe with time.

Basidiospores [30/2/2] (7.8) 9.5–12.8 (13.31)  (5.7) 6.3–7.8 (8.9), Q = (1.21) 1.39–1.91 (1.98), avQ = 1.62, elliptical, smooth, uniguttulate, brownish in 5% KOH. Hymenal cystidia (18.5) 19.2–21.75 (25.3)  (7.5) 7.7–8.6 (9.1) µm, clavate, thin walled, hyaline in 5% KOH. Basidia (14.8) 26.4–31.2 (36.7)  (10.08) 10.5–10.83 (11.8) µm, clavate , thin walled, guttulate, hyaline in 5% KOH. Stipitipellis (4.5) 4.7–8.7 (9.16) µm wide, filamentous, rarely branched, septate hyphae, clamp connections not observed, hyaline in 5% KOH. Pileipellis (3.8) 4.9–6.3 (6.6) µm wide, filamentous, fusoid terminals, frequently septate, clamped septa common, hyaline in 5% KOH.

Material examined: Pakistan, Khyber Pakhtunkhwa province, Hazara division, Mansehra district, Kaghan valley (Khanian), 2500 m asl, on soil under Cedrus deodara, 3 Aug 2014, Muhammad Burhan Ino; SJ103(LAH35042); Malakand division, Swat district, Mashkun, 2500 m asl, on soil under C. deodara, 5 Sep 2013, Sana Jabeen MTI; (LAH35043) .

Molecular phylogenetic characterization (Figure 50)

Sequencing of the PCR products of nrITS region of Inocybe flavellorimosa yielded 798–817 base pairs by using ITS1F and ITS4 primers. A consensus sequence of 631 base pairs was subjected to BLAST. It was found 89% identical to I. rimosa (Bull.) P. Kumm. (JF908172) I. var. rimosa (HQ604622) and I. bulbosissima (Kühner) Bon

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(FJ904159, JX436913, AM882765 & FJ904160) with 91–92% query cover and 0.0 E value.

To reconstruct phylogeny, closely related ITS sequences were retrieved from the GenBank. Auritella foveata C.K. Pradeep & Matheny (GU062740) was chosen as out group (Larsson et al., 2009). Sequences of Inocybe flavellorimosa clustered with I. bulbosissma, I. rimosa, I. melliolens Kühner , I. sororia Kauffman and I. umbrinella Bres. in a same clade within sub genus Rimosae sensu stricto following classification by Larsson et al. (2009). It gets separated from all these taxa with 95% bootstrap value.

Comments

Inocybe flavellorimosa nom. prov. is characterized by its yellowish brown cap with dark brown striations, highly rimose margins and stipe giving off its stipitipellis with time and a rimose stipe base. In ITS data set analysis it clustered in sub genus Rimosae. Taxa within this group were separated into A–F clades. Inocybe flavellorimosa clustered in clade A. This clade corresponds to I. rimosa in a wide sense and includes sequences mostly from England, Estonia, France, and Scandinavia. They represent most of the large variation in macro-morphological characters demonstrated by the many varieties described (Heim, 1931; Favre, 1955; Kühner, 1988; Bon, 1997). specimens show a great variation in cap colors from pale to ochraceous yellow brown to dark brown. Inocybe flavellorimosa looks more or less similar to a typical I. rimosa, but it showed a peeling off stipitipellis and highly rimose cap and the base of stipe. Anatomically these both taxa are almost similar. Inocybe melliolens originated from France, also morphologically looks like a typical I. rimosa (Bon, 1997).

Inocybe sororia specimens sampled for phylogenetic analysis includes Swedish material and which they deviate from the descriptions given by Kauffman (1924) and Stuntz (1947). These descriptions are found uncertain about the identity of Swedish collections. Inocybe bulbosissima includes specimens usually regarded as a variety of I. rimosa and then named I. fastigiata var. alpina. Inocybe umbrinella exhibit warm yellowish to reddish brown caps with a dark centre and contrasting strongly rimose and lighter periphery while I. flavellorimosa bears yellowish cap with brown striations, but the micro-morphology is almost identical to I. rimosa. All theses taxa formed their separate lineages within clade A of subgenus Rimosae with a strong boot strap value.

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Figure 48. Morphology of Inocybe flavellorimosa. A & B. Basidiomata LAH35042 (holotype). Bars: A & B = 1 cm.

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Figure 49. Anatomy of Inocybe flavellorimosa. A–E. LAH35042. A. Basidiospores; B. Hymenal cystidia; C. Basidia and basidiole; D. Stipitipellis; E. Pileipellis. Bars: A–E = 10 µm.

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58 I alba SJ146 49 100 I alba SJ102 I alba SJ147 84 I alba SJ102b HQ604626 Inocybe sororia 22 93 KP783443 Inocybe sororia JQ408750 I breviterincarna JQ408753 I breviterincarna E 100 JQ408754 I breviterincarna 72 JQ408751 I breviterincarna JX630909 I dulcamaroides FJ904127 Inocybe dulcamaroides 28 FJ904126 Inocybe dulcamaroides 100 FJ904127 I dulcamaroides FJ904126 I dulcamaroides 90 FJ904134 I arenicola FJ904133 I arenicola FJ904124 I mimica 82 I mimica SJ21b I mimica SJ111 I mimica SJ154 100 KJ726737 I mimica D 41 I mimica SJ21 51 KJ546158 I mimica KF056319 I mimica I mimica SJ73 I mimica SJ27 KJ700456 I mimica 99 JF908260 I spuria FJ904139 I spuria JQ408794 I spuria 88 JF908162 I squamata 100 AM882780 I squamata 53 97 FJ904132 I squamata 92 FJ904136 I squamata 43 FJ904137 I hygrophorus C JX436912 I flavella 64 25 FJ904128 I cf flavella JQ724027 I flavella Rimosae 96 JQ724026 I flavella JQ724025 I flavella 61 FJ904129 Inocybe cf flavella FJ904129 I cf flavella JQ408789 I sp 100 AM882769 I obsoleta 78 AM882770 I obsoleta B 57 AM882772 I perlata 100 AM882771 I perlata 100 I flavellorimosa SJ103b I flavellorimosa SJ103 33 FJ904166 I umbrinella 98 66 FJ904163 I umbrinella FJ904165 I umbrinella KR733590 I cf rimosa 95 56 FJ904158 I bulbosissima 59 FJ904159 I bulbosissima 21 FJ904160 I bulbosissima 48 AM882765 I bulbosissima 45 86 AM882777 Inocybe rimosa HQ215781 Inocybe sp HQ604622 Inocybe rimosa var rimo JF908172 Inocybe rimosa 49 AM882767 Inocybe rimosa A 29 39 JQ408765 Inocybe sp I sp15 SJ25 FJ904153 Inocybe cf rimosa FJ904147 Inocybe rimosa 23 AM882763 I rimosa 56 AM882761 I rimosa 98 AM882762 I rimosa FJ904148 I melliolens 41 FJ904149 I melliolens AM882844 I rimosa HQ604610 I sororia HQ604618 I sororia 100 HQ604617 I sororia 69 HQ604607 I sororia GU062740 Auritella foveata Auritella

0.05 Figure 50. Molecular phylogenetic analysis of Inocybe spp. based on ITS sequences. The evolutionary history was inferred by the Maximum Likelihood method using General Time Reversible model. The analysis involved 78 nucleotide sequences. There were a total of 796 positions in the final dataset. Sequences generated during this study are marked with .

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Inocybe kohistanensis Jabeen, Ahmad & Khalid, Turk. J. Bot. 47: 231 (2016)

Jabeen et al., 2016 (Annexure)

Inocybe mimica Massee, Ann. Bot., Lond. 18: 492 (1904) (Figures 51 & 52)

Basidiomata medium. Pileus upto 4.5 cm wide, spherical, conical becoming wide towards maturity, umbo prominent brown (2.5YR4/8) becoming light brown 5YR6/10 to yellowish brown (10YR7/8) towards the margins, surface dry, fibrillose, margins incurved, rimose when mature. Lamellae regular, adnexed, edges even to slightly fimbriate, moderately close, up to 0.3 cm broad, cream (2.5Y9/2) to yellowish (10YR8/6) or brownish (10YR6/6) becoming golden brown (10YR7/10) when dry. Lamellulae versatile in length. Stipe 6.5 × 1.0 cm, base swollen up to 2 cm wide, becoming narrower towards the apex, off white context with brownish (10YR78) patches, central, cylindrical, surface granular to fibrillose towards the base.

Basidiospores [50/5/5] (9.3) 10.3–13.2 (14.5)  (6.1) 6.4–7.2 (8.1), Q = (1.21) 1.42–1.70 (1.88), avQ = 1.39, smooth, ellipsoid, apiculus prominent, brownish in 5% KOH. Basidia (23.5) 25.2–35.4 (40.4)  (9.2) 9.4–114.0 (16.5) µm, wall up to 3 µm, clavate, densely guttulate, 4-spored, hyaline in 5% KOH. Hymenalcystidia (30.7) 56.9– 120.3 (130.3)  (20.8) 21.4–28.2 (30.9) µm, wall up to 3 µm, crystalliferous apex also observed, hyaline in 5% KOH. Caulocystida 55.3–70.5  9.4–16.6 µm, cylindrical to clavate or fusoid, hyaline in 5% KOH. Stipitipellis filamentous, (2.1) 2.6–7 (9.7) µm wide, rarely branched, rarely septate hyphae. Pileipellis filamentous, hyphae (3.3) 4.5–8.5 (18.2) µm wide, clavate to fusiform terminals, hyaline in 5% KOH

Material examined: Pakistan, Khyber Pakhtunkhwa province, Malakand division, Swat district, Kalam, 2400 m asl, on soil under Cedrus deodara, 4 Sep 2013, Sana Jabeen & Aamna Ishaq K2-6; SJ73(LAH35048); Muhammad Abdur Rehman K2-24; SJ153 (LAH35051); Sana Jabeen & Abdul Nasir Khalid K5; (LAH35052); K3-7; SJ154 (LAH35053); Mashkun, 2500 m asl, on soil under C. deodara, 5 Sep 2013, Sana Jabeen & abdul Nasir Khalid K4-6; SJ21 (LAH35050); Abdul Nasir Khalid MT2C-SA; SJ27 (LAH35049).

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Molecular phylogenetic characterization (Figure 53)

Sequencing of the PCR products of nrITS region of Inocybe mimica yielded 734– 881 base pairs by using ITS1F and ITS4 primers. Consensus sequence of 654 base pairs was subjected to BLAST at NCBI. It showed 99% identity to I. mimica sequences from India ( KF056319), Pakistan (KJ726737, KJ546158 & KJ700456) and Sweden (AM88278 & FJ904124) with 99–100% query cover and 0.0 E value.

For phylogenetic analysis, closely related ITS sequences were retrieved from the GenBank. Auritella foveata C.K. Pradeep & Matheny (GU062740) was chosen as out group (Larsson et al., 2009). Sequences of Inocybe mimica generated during this study clustered with similar taxa from India, Pakistan and Sweden in a same clade within sub genus Rimosae sensu stricto following classification by Larsson et al. (2009).

Comments

Inocybe mimica was first described by Massee (1904) from Europe. Records have been found from India, Itlay, Pakistan and Sweden (Larsson et al., 2009; Bizio & Ferisin, 2013; Saba et al., 2015). The species is closely related to I. arenicola and clusterd within clade F within sub genus Rimosae. Although, the taxa belonging to this group are more or less similar in appreance, intera specific morphological variations has also been observed. Both of these taxa are found similar by their comparatively larger smooth spores (Larsson et al., 2009). The species is reported as rare from Europe (Larsson et al., 2009; Bizio & Ferisin, 2013), but it has been found abundently from different areas in Swat, Pakistan during this study.

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A B C

D E F

G H Figure 51. Morphology of Inocybe mimica. A–H. Basidiomata. A & F. LAH35042; B & C. LAH35043; D & E. LAH35045; G. LAH35046; H. LAH35047. Bars: A–H = 1 cm.

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A B

C D

E F

Figure 52. Anatomy of Inocybe mimica. A–F LAH35042. A. Basidia; B. Basidiospores; C. Cheilocystidia; D. Plurocystidia; E. Stipitipellis; F. Pileipellis. Bars: A–F = 10 µm.

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57 KF056319_I_mimica FJ904124_I_mimica KJ726737_I_mimica KJ546158_I_mimica 53 I_mimica_SJ27 I_mimica_SJ73 KJ700456_I_mimica 98 I_mimica_SJ111 I_mimica_SJ21 F 90 46 I_mimica SJ121b I mimica SJ154 43 FJ904134_I_arenicola 98 FJ904133_I_arenicola JQ408750_I_breviterincarna 16 JQ408753_I_breviterincarna 100 JQ408754_I_breviterincarna 72 JQ408751_I_breviterincarna 18 JQ408776_I_cf_rimosa E 100 JQ408775_I_cf_rimosa FJ904127_I_dulcamaroides FJ904126_I_dulcamaroides D 71 100 JX630909_I_dulcamaroides_ 99 JQ408794_I_spuria JF908260_I_spuria FJ904139_I_spuria 94 JF908162_I_squamata 99 96 AM882780_I_squamata FJ904132_I_squamata 88 FJ904136_I_squamata 61 C 49 FJ904137_I_hygrophorus Rimosae JX436912_I_flavella 77 JQ724026_I_flavella 36 98 FJ904128_I_cf_flavella_ JQ724027_I_flavella 57 JQ724025_I_flavella FJ904129_I_cf_flavella_ JQ408789_I_sp 100 AM882769_I_obsoleta 61 AM882770_I_obsoleta B 84 AM882772_I_perlata 100 AM882771_I_perlata 72 FJ904165_I_umbrinella 99 FJ904163_I_umbrinella FJ904166_I_umbrinella 97 KR733590_I_cf_rimosa_ FJ904160_I_bulbosissima 17 AM882765_I_bulbosissima 43 FJ904159_I_bulbosissima 12 FJ904158_I_bulbosissima 83 HQ604617_I_sororia 8 HQ604607_I_sororia A 91 HQ604618_I_sororia 16 HQ604610_I_sororia HQ604611_I_sororia 61 97 FJ904148_I_melliolens FJ904149_I_melliolens AM882763_I_rimosa 49 AM882762_I_rimosa 4 AM882844_I_rimosa 17 AM882761_I_rimosa GU062740_Auritella_foveata Auritella

0.05 Figure 53. Molecular phylogenetic analysis of Inocybe mimica based on ITS sequences. The evolutionary history was inferred by the Maximum Likelihood method using General Time Reversible model. The analysis involved 61 nucleotide sequences. There were a total of 777 positions in the final dataset. Sequences generated during this study are marked with .

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Inocybe oblectabilis (Britzelm.) Sacc., Syll. fung. (Abellini) 11: 54 (1895)

(Figures 54 & 55)

Basidiomata medium. Pileus up to 3.5 cm wide, spherical, conical becoming wide towards the margins, pileipellis golden brown (5YR6/12) having dark brown (5YR4/8) striations, umbo prominent, blackish brown (5YR2/4) becoming lighter towards the margins, surface dry, fibrillose, margins incurved, rimose when mature. Lamellae regular, adnexed, edges slightly fimbriate, moderately close, up to 0.3 cm broad, grayish (7.5YRB/2) when young becoming golden brown (5Y6/12) when mature and dry. Lamellulae versatile in length. Stipe 7.2 × 1.1 cm, base marginately swollen up to 2 cm wide, becoming narrower towards the apex, surface pruinose to smooth or fibrillose.

Basidiospores [30/3/3] (8.7) 12.3–15.2 (17.4)  (7.5) 8.7–10.2 (11.2) µm, Q = (1.10) 1.25–1.55 (1.72), avQ = 1.43, elongated, angular, nodulose, with 8–9 nodules, yellowish 5% KOH, guttulate. Basidia (32.5) 38.2–40.4 (43.3)  (15.0) 15.6–16.8 (20.5) µm, clavate, thin walled, 4-spored, densely guttulate, hyaline in 5%KOH. Cheilocystidia (61.2) 64.7–79.9 (87.5)  (24.5) 28.1–32.2 (33.5) µm, with crystalliferous apex, pale yellow to hyaline in 5% KOH. Pleurocystidia (50.7) 63.9–76 (80.2)  (20.8) 21–28 (31.9) µm, densely crystalliferous apex, pale yellow to hyaline in 5% KOH. Stipitipellis filamentous, (3.8) 4.9–8.5 (13.7) µm wide, rarely branched, pale yellow, septate hyphae, clamp connections observed. Pileipellis filamentous, hyphae (3.3) 4.4–8.7 (12.2) µm wide, branched, fusiform terminals, yellowish brown, clamp connections frequent.

Material examined: Pakistan, Khyber Pakhtunkhwa province, Malakand division, Swat district, Kalam, 2400 m asl, on soil under Cedrus deodara, 4 Sep 2013, Abdul Nasir Khalid K2-12; SJ14 (LAH35054); Sana Jabeen K3-4; SJ47 (LAH35055); K3-5; SJ80 (LAH35056); associated with C. deodara, 4 Sep 2013, Sana Jabeen & Abdul Nasir Khalid KF9; SA214 (LAH-EM19-2013); KF11; SA342 (LAH-EM18-2013).

Molecular phylogenetic characterization (Figure 56)

Sequencing of the PCR products of nrITS region of Inocybe oblectabilis yielded 655–728 base pairs by using ITS1F and ITS4 primers. A consensus sequence of 584 base pairs was subjected to BLAST at NCBI. It showed 96–98% identity to I. pallida Velen.

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(JF908198) and I. oblectabilis (Britzelm.) Sacc. (JF908204, AM882831, AM882932 & AM882833) from Sweden and USA 97–100% query cover and 0.0 E value.

For phylogenetic analysis, closely related ITS sequences were retrieved from the GenBank. Taxa from sections Rimosae and Inosperma were selected as out groups. Sequences of Inocybe oblectabilis generated during this study clustered with similar taxa in same clade within section Marginatae sub section Oblectabilies with strong bootstrap value.

Comments

Inocybe oblectabilis was first identified as Agaricus oblectabilis Britzelm in 1890 from Europe (Britzelmayr, 1890). The species has been recorded from Austria, Canada, Estonia, Norway and Sweden (http://www.gbif.org/species/2527973). It has also been reported from Finland (Kobayashi, 2003) and Pakistan (Ahmad et al., 1997). A recent detailed morphological data is given by Kobayashi in 2003 based on Japanese collection. Morphological characters of I. oblectabilis from Pakistan coincide with those reported from Europe (Britzelmayr, 1890; Kühner, 1933; Stangl, 1989) and Japan (Kobayashi, 2003). Molecular data based on ITS sequences also support its identification with I. oblectabilis from Sweden. It clustered within section Marginatae subsection Oblectabilies which is separated by the sub section Praetervisae based on more or less pinkish to reddish coloration on the stipe, at least, on the upper surface of the stipe following the pattern proposed by Bon (1988). Its ectomycorrhizal association is being reported for the first time during this study.

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Figure 54. Morphology of Inocybe oblectabilis. A–F. Basidiomata. A & D. LAH35055; B & C. LAH35054; E & F. LAH35056. Bars: A–C = 1.3 cm, D–F = 1.4 cm.

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A B

C

D

E F Figure 55. Anatomy of Inocybe oblectabilis. A–F. LAH35054. A. Basidiospores; B. Basidia; C. Stipitipellis; D. Cheilocystidia; E. Pileipellis; F. Pleurocystidia. Bars: A–F = 10 µm.

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54 KJ938773 I xanthomelas 61 KJ938774 I xanthomelas 100 HQ586858 I straminipes KJ938775 I xanthomelas 99 68 HQ586868 I straminipes KJ938770 I krieglsteineri KJ938771 I krieglsteineri 81 100 KJ938772 I krieglsteineri KJ938768 I krieglsteineri FJ755798 I saliceticola 46 AM882717 I saliceticola 100 FJ755796 I saliceticola isolate 71 FJ755799 I saliceticola 49 HQ604598 I praetervisa HQ604599 I praetervisa AM882723 I hirculus 100 Praetervisae 91 JX436916 I hirculus 99 JF908234 I margaritispora 82 KJ938781 I flavobrunnescens KJ938782 I flavobrunne 100 JF908120 I margaritispora 37 KJ938784 I flavobrunnescens KJ938769 I krieglsteineri KJ938783 I flavobrunnescens 100 AM882724 I salicis Marginatae 56 FJ531872 I hirculus 81 KJ938778 I ochracea 67 100 FJ755803 I ochracea KJ938777 I ochracea 35 EU523561 I intricata HQ586869 I obtusiuscula 67 100 EF655704 I rufofusca AJ889951 I asterospora 100 AJ889950 I asterospora 20 AJ889951 I asterospora HQ604559 I napipes 100 HQ604556 I napipes 74 KT897911 I kohistanensis 100 KP316244 I kohistanensis KP316245 I kohistanensis KP316243 I kohistanensis 72 Oblectabiles 68 I dunensis JF908262 95 100 I dunensis JF908093 I dunensis JF908092 JF908198 I pallida 89 99 I oblectabilis M882833 38 I oblectabilis AM882832 100 I oblectabilis SJ342 I oblectabilis SJ214 6887 I oblectabalis SJ14 100 FJ904159 I bulbosissima Rimosae FJ904144 I rimosa FJ904174 I quietiodor Inosperma 99 AM882955 I cookei

0.05 Figure 56. Molecular phylogenetic analysis of Inocybe oblectabilis based on ITS sequences. The evolutionary history was inferred by the Maximum Likelihood method using General Time Reversible model. The analysis involved 54 nucleotide sequences. There were a total of 737 positions in the final dataset. Sequences generated during present investigation are marked with .

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Genus Rhizopogon

Rhizopogon flavus nom. prov. (Figure 57)

Etymology: The specific epithet refers to the yellowish color of the exoperidium.

Basidiomata sub globose, yellowish (5Y8/6) to brown (2.5Y6/8), 2.5–4.0 cm high and 2.0–4.0 cm wide, semi hypogeous, pale yellow when fresh, turning brown with age, covered with thin membrane of hyphae, no change in color when bruised. Rhizomorphs up to 1.8 cm long, frequent at the base, well developed, whitish, branched, encrusted with debris and soil particles, dehiscence not observed. Exoperidium pale yellowish, persistent. Gleba off white to yellow or brownish, firmly spongy.

Basidiospores [20/2/1] 4.6–8.0 × 3.0–4.0 µm, ellipsoid to oblong, smooth, pale green in 5% KOH, non amyloid. Gleba bears hyaline to brown, thin walled, branched, and septate hyphae.

Material examined: Pakistan, Khyber Pakhtunkhwa province, Malakand division, Swat district, Mashkun, 2400 m asl, semi hypogeous in soil, in groups under Cedrus deodara, 3 Sep 2013, Sana Jabeen K4-10; SJ39 (LAH35057).

Molecular phylogenetic characterization (Figure 58)

Sequencing of the PCR products of nrITS region of Rhizopogon flavus yielded 733–751 base pairs by using ITS1F and ITS4 primers. A consensus sequence of 643 base pairs was subjected to BLAST at NCBI. It showed 92% identity to R. roseolus (Corda) Th. Fr. from Slovenia (AJ810047, AJ810051 & AJ810052) and Spain (AJ810057, AJ810062–AJ810064 ) with 100% query cover and 0.0 E value.

For phylogenetic analysis, closely related ITS sequences were retrieved from the GenBank. Suillus sibiricus (Singer) Singer 1945 (KM882916) was chosen as out group. Rhizopogon flavus clustered in a same clade with R. rubescens (Tul. & C. Tul.) Tul. & C. Tul. and R. roseolus with strong boot strap support. It gets separated from R. rubescens forming a sister lineage of its own.

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Comments

Rhizopogon flavus nom. prov. is characterized by its yellow exoperidium, off white to brownish gleba and smooth ellipsoid spores. It does not show any change in color upon bruising. The species can be compared with its closely related taxa. It can be distinguished from R. rubescens and R. roseolus on the basis of color of exoperidium. Rhizopogon rubescens and R. roseolus bear reddish to pink patches on the exoperidium, while R. flavus lacks such reddish or pinkish coloration. Moreover, the glebal part of R. flavus does not show any color change upon bruising, while R. rubescens shows pink glabal coloration upon incision (Baseia & Milanez, 2002).

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A

B C Figure 57. Morphology and anatomy of Rhizopogon flavus. A–C. LAH35003. A. Basidiomata; B. Basidiospores; C. Hyphae from gleba. Bars: A = 1cm; B & C = 6 µm.

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DQ179127_Rhizopogon_roseolus AJ810053_Rhizopogon_roseolus 100 AJ810047_Rhizopogon_roseolus AJ810052_Rhizopogon_roseolus 90 AJ810051_Rhizopogon_roseolus 50 AJ810065_Rhizopogon_roseolus AJ810067_Rhizopogon_roseolus 68 AJ810064_Rhizopogon_roseolus 92 AJ810063_Rhizopogon_roseolus AJ810062_Rhizopogon_roseolus JF908636_Rhizopogon_vulgaris KF482486_Rhizopogon_vulgaris 40 AJ810074_Rhizopogon_roseolus 82 26 74 AJ810056_Rhizopogon_roseolus 30 AJ419209_Rhizopogon_roseolus 40 AJ810057_Rhizopogon_roseolus 64 Rhizopogon flavus SJ39b 100 Rhizopogon flavus SJ39 Rhizopogon flavus F39b AJ966744_Rhizopogon_roseolus 58 JX907816_Rhizopogon_rubescens 100 DQ068965_Rhizopogon_rubescens AJ277644_Rhizopogon_rubescens AM085529_Rhizopogon_sardous AJ810071_Rhizopogon_roseolus 42 NR_119470_Rhizopogon_graveolens 84 AJ419211_Rhizopogon_roseolus EU379678_Rhizopogon_roseolus 96 HM036649_Rhizopogon_roseolus AB685401_Rhizopogon_roseolus 30 KF990475_Rhizopogon_roseolus 40 AB505222_Rhizopogon_sp 50 AB685403_Rhizopogon_roseolus KC152205_Rhizopogon_pseudoroseol 100 KC152206_Rhizopogon_pseudoroseol KC152207_Rhizopogon_pseudoroseol 88 AF266710_Rhizopogon_sp 76 KP859266_Rhizopogon_abietis KP859267_Rhizopogon_abietis 64 AF062934_Rhizopogon_vulgaris 100 GQ267482_Rhizopogon_luteorubesce 74 NR_119471_Rhizopogon_luteorubesc KM882916_Suillus_sibiricus Out group

0.01 Figure 58. Molecular phylogenetic analysis of Rhizopogon flavus based on ITS sequences. The evolutionary history was inferred by the Maximum Likelihood method using Jukes-Cantor model. The analysis involved 43 nucleotide sequences. There were a total of 682 positions in the final dataset. Sequences generated during present investigation are marked with .

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Genus Russula

Russula anthracina Romagn., Bull. mens. Soc. linn. Lyon. 31: 173 (1962)

Jabeen et al., 2016 (Annexure)

Russula amethystina Quél., Compt. Rend. Assoc. Franç. Avancem. Sci. 26: 450 (1898)

(Figures 59 & 60)

Pileus 2–5.2 cm broad, campanulate when young becoming convex to infundibuliform at maturity; purple 2.5R2/6 to brownish purple (5RY/6) or reddish violet (5R4/12), sometimes cream (2.5Y8/6) at the edges; surface smooth, dry; margins smooth, incurved when young, becomming straight to uplifted at maturity. Lamellae up to 0.35 cm broad, creamy white (2.5Y9/2); regular, adnexed, subdistant to close, edges entire. Lamellulae few. Stipe 3–7 cm × 0.7–1.9 cm, central, clavate, surface fibrillose, bare, white to cream (5Y9/2).

Basidiospores [60/6/6] (8.2) 8.7–9.9 (10.4) × (6.0) 7.3–8.2 (8.7) µm, Q = (1.0) 1.05–1.35 (1.41), avQ = 1.21; globose to subglobose, ornamented, prominent warts accompanied with some low warts, apiculus prominent, pale yellow in 5% KOH, amyloid in Malzer‟s reagent. Basidia (41.3) 46.6–48.9 (52.1) × (9.7) 10.6–12.7 (12.6) µm, clavate, guttulate, thin walled, wall up to 3 µm thick, pale yellow to hyaline in 5% KOH. Cheilocystidia (43.2) 49.6–57.3 (71.1) × (7.4) 8.9–10.1 (11.5) µm, fusiform, mucronate to rostrate, guttulate, pale yellow to hyaline in 5% KOH. Pleurocystidia (57.3) 59.1–68.2 (72.7) × (7.2) 7.8–9.0 (11.8) µm, variable in shape, from fusiform, mucronate to rostrate, pale yellow to hyaline in 5% KOH. Pileipellis hyphae (5.45) 8.18–11.7 (15.4) µm wide, filamentous, branched, pale yellow to hyaline in 5% KOH, septa infrequent, clamp connections and pileocystidia not observed. Stipitipellis hyphae (6.36) 8.18–12.7 (14.5), filamentous, branched, pale yellow to hyaline, in 5% KOH, rarely septate, clamp connections not observed.

Material examined: Pakistan, Khyber Pakhtunkhwa province, Malakand division, Swat district, Kalam, 2400 m asl, on soil under Cedrus deodara, 3 Sep 2013, Sana Jabeen & Aamna Ishaq K7; SJ18 (LAH35057); 4 Sep 2013, Sana Jabeen, Abdul Rehman Khan Niazi & Abdul Nasir Khalid K2-11; SJ29 (LAH35058); Mashkun, 2500 m asl, on soil under C. deodara, 5 Sep 2013, Sana Jabeen, Ishtiaq Ahmad & Abdul Nasir Khalid K4-27;

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SJ25 (LAH35059); Sana Jabeen & Abdul Nasir Khalid K4-12; SJ84 (LAH35060); K4- 28; SJ31 (LAH35061); K4-29; SJ22 (LAH35062); MT2-E; SJ92 (LAH35063); MT2-1 (LAH35064).

Morphological characterization of ectomycorrhiza (Figure 61)

Ectomycorrhizal system irregular to monopodial pinate or monopodial pyramidal; up to 1.8 cm long; tips 0.2–1 cm long and up to 1.2–1.7 mm wide, apex 0.2 mm wide, base up to 0.23 mm wide; tips yellowish brown (5YR5/10) to dark brown (2.5YR2/4) or black at maturity; unramified ends straight to curved; Mantle pale yellow (10YR8/8); soil particles adherent to the mantle surfae; host tissue visible under the mantle surface.

Rhizomorphs infrequent, pale yellow (5YR7/10) to hyaline. Emanating hyphae frequent, branched, pale yellow (5YR7/10) to hyaline.

Mantle plectenchymatous in all layers (mantle type E, Agerer, 1987–2002; Agerer & Rambold, 1998), non-gelatinous, hyphae thin long, branched, septate, septa infrequent, hyphae membranaceously and plasmatically pale yellow hyaline, upto 2.3 µm wide, contents clear. Emanating hyphae straight, elongated, cylindrical, infrequently septate, 2.7 µm in diameter, clamp connections absent, surface smooth, contents clear, ramification frequent, Y-shaped; hyphal endings simple, anastomosing without septum. Cystidia infrequent, bottle shaped, upto 12.3 µm in length, inflated base, upto 4.79 µm wide, a long straight neck 1.8 µm in diameter. Rhizomorphs hyphae branched, uniform and loosely arranged, intermingled, 2.3 µm in diameter, septate, clamp connections not observed, pale yellow to transparent.

Material examined: Pakistan, Khyber Pakhtunkhwa province, Malakand division, Swat district, Kalam, 2400 m asl, associated with Cedrus deodara, 24 Aug 2014, Sana Jabeen & Abdul Rehman Khan Niazi KF6b2; SA217 (LAH-EM47-2013).

Molecular phylogenetic characterization (Figure 62)

Sequencing of the PCR products from the DNA of Russula amethystina using ITS1F and ITS4 primers yielded 722–847 base pairs. A 625 base pair consensus sequence was obtained by trimming the motifs. BLAST searched revealed its 98–99% identity with R. amethystina (JF908682) from Italy, R. gilva Zvára (KF002762) from China, R. lutea

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(Huds.) Gray (HQ604848 & KF679818) from Pakistan and R. postiana Romell (KF850410 & AF230898) from Spain. with 98–100% query cover and 0.0 E value.

Published sequences from different Russula sections were included to reconstruct a phylogeny with Boletus reticuloceps (Zang et al.) Wang & Yao (EU231968) as outgroup. The analysis revealed four major clades with strong bootstrap support. Russula amethystina clustered with other related taxa in section Tenellae with strong boost strap support.

Comments

Russula amethystina has been named after its amethyst colored pileaus surface. The taxon is morphologically releted to R. azurea Bres. and R. turci Bres. It is very hard to distiguish these taxa on the basis of macro morphological features. ITS sequence comparisons separate these taxa in distinct clades within the same section. Russula amethystina is an europuan taxon. It has been reported from Italy (Osmundson et al., 2013) and Slovakia (Caboň, 2013). Records have also been found from Austria, Czech Republic, France, Germany, Morocco, Norway, Romania, Russia, Spain and Sweden (http://www.gbif.org/). Sequences generated from local fruit bodies and ectomycorrhizal tissue clustered with each other along with R. amethystina from Italy. Its collections during this study represent an addition to the mycobiota of Pakistan.

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Figure 59. Morphology of Russula amythistina. A–L. Basidiomata. A. LAH35058; B. LAH35059; C. LAH35060; D. LAH35057; E & F. LAH35061; G & H. LAH35063; I & J. LAH35064; K & L. LAH35062. Bars: A–L = 1.3 cm.

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Figure 60. Anatomy of Russula amythystina. A–F. LAH35058. A. Basidiospores; B. Basidia; C. Pleurocystidia; D. Cheilocystidia; E. Pileipellis; F. Stipitipellis. Bars: A = 5.4 µm; B = 11 µm, C = 12.3 µm; D = 11.2 µm; E & F = 21.1 µm.

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Figure 61. Ectomycorrhiza of Russula amythestina. A–G. LAH-EM12-2014. A & B. Ectomycorrhizal morphotypes; C. Cystidia from outer mantle surface; D. Inner mantle layer; E. Outer mantle layer; F. Emanating hyphae; G. Rhizomorphs. Bars: A & B 0.3 cm, C = 4.6 µm; D & E = 6 µm; F = 8.1 µm; G = 7 µm.

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KT953617 Russula amethystina SJ217 KT953616 Russula amethystina SJ92 KT953614 Russula amethystina SJ59 KT953612 Russula amethystina SJ18 95 KF679818 Russula lutea KT953613 Russula amethystina SJ29 54 KT953615 Russula amethystina SJ84 JF908682 Russula amethystina AF230898 Russula postiana KF850410 Russula postiana 95 77 KF002762 Russula gilva DQ422022 Russula risigallina AY061713 Russula risigallina 99 83 JF908702 Russula velenovskyi Tenellae 54 HQ604848 Russula lutea 76 AJ971402 Russula paludosa 99 JF908659 Russula paludosa 44 JQ888199 Russula paludosa AF418627 Russula solaris 99 91 GU234063 Russula chamiteae 50 FJ845432 Russula decolorans 50 AY245542 Russula californiensis AF418631 Russula firmula 28 97 AF418630 Russula veternosa HQ604850 Russula murrillii AY061716 Russula roseipes 99 54 88 JQ711999 35 EU819426 Russula mariae 37 EU819437 AF418612 55 Heterophyllae 52 AF418611 Russula parazurea AF418610 Russula vesca 99 AF418609 Russula heterophylla 48 GU371295 Russula livescens 99 50 AY061700 Russula insignis JQ622344 Russula pulverulenta 85 AB211275 Russula sororia 99 JQ622327 Russula amoenolens 99 JN681168 Russula cerolens AJ438037 Gymnomyces ammophilus 85 Ingratae 14 HE647707 Russula 23 KF002757 Russula subfoetens 60 HQ677769 Russula illota 36 KF245527 Russula laurocerasi 13 KF245530 Russula foetens AY239335 Gymnomyces parksii 81 61 KC152107 Gymnomyces fallax 50 EU598197 Russula eccentric 63 DQ422027 Russula polyphylla KF306038 Russula cantharellicola 68 AB291746 Russula subnigricans KC581314 91 JF834355 Russula albonigra 59 AB291765 Compactae 83 75 EU303008 Russula dissimulans 47 JQ888194 Russula adusta 72 JF834370 Russula aff acrifolia JF908673 Russula anthracina 94 EU526006 Russula cascadensis 47 65 JF834363 Russula acrifolia EU231968 Boletus reticuloceps Out group

0.05 Figure 62. Molecular phylogenetic analysis of Russula amethystina based on ITS sequences. The evolutionary history was inferred by the Maximum Likelihood method based Jukes-Cantor model. The analysis involved 61 nucleotide sequences. There were a total of 835 positions in the final dataset. Sequences generated from during this study are marked with .

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Russula delica Fr., Epicr. syst. mycol. (Upsaliae): 350 (1838) (Figures 63 & 64)

Pileus upto 10 cm broad, depressed from the middle, margins incurved when young, becomming infundibuliform when mature. Surface range white with light brown (10YR7/8) patches in the centre; surface dry and smooth. Lamellae up to 0.3 cm broad, white becoming brownish (10YR8/8) with age, regular, adnexed, close, edges entire. Lamellulae frequent, variable in length. Stipe 4.7 × 2 cm, white, central, cylindrical, slightly narrower towards the base. Surface smooth to finely wrinkled.

Basidiospores [50/5/5] (8.2) 9.9–10.5 (12.3) × (8.0) 8.5–9.7 (10.5) µm, Q = (1) 1.05–1.4 (1.22), avQ = 1.13 ± 0.04, globose to subglobose, echinulate, apiculus prominent, pale yellow in 5% KOH, amyloid in Melzer‟s reagent. Basidia (60.4) 64.5– 69.6 (72.4) × (13.5) 14.2–15.4 (16.7) µm, clavate, wall up to 3 µm, pale yellow in 5% KOH. Pleurocystidia (75.9) 78.2–96.6 (104.7) × (10.5) 12.8–17.5 (19.6) µm, fusoid, guttulate, hyaline in 5% KOH. Cheilocystidia (62.4) 68.3–75.6 (85.7) × (10.4) 12.2–15.6 (17.8) µm, fusiform, guttulate, hyaline in 5% KOH. Pileipellis hyphae 12.8–31.5 µm wide, filamentous, branched, hyaline in 5% KOH, septate, clamp connections not observed. Stipitipellis hyphae 15.1–30.2 µm wide, filamentous, branched, hyaline in 5%KOH, septate, clamp connections not observed.

Material examined: Pakistan, Khyber Pakhtunkhwa province, Malakand division, Swat district, Kalam, 2400 m asl, on soil under Cedrus deodara, 3 Sep 2013, Sana Jabeen K4; SJ50 (LAH35065); 4 Sep 2013, Abdul Nasir Khalid K2-13; (LAH35066); Mashkun, 2500 m asl, on soil under C. deodara, 5 Sep 2013, Sana Jabeen K4-25; (LAH35067); Punjab province, Rawalpindi division & district, Kuza Gali, 2500 m asl, on soil under C. deodara, 21 Sep 2013, Sana Jabeen KG-1; SJ51 (LAH35068); KG-2; SJ89 (LAH35069).

Molecular phylogenetic characterization (Figure 65)

Sequencing of the PCR products from Russula delica using ITS1F and ITS4 primers yielded 688–790 base pairs. A consensus sequence of 623 base pairs was obtained by trimming the motifs. BLAST revealed that it was 96–98% identical to R. delica sequences (KF432955 & AF418605) from Belgium with 98% query cover and 0.0 E value.

Published sequences from different Russula sections were included to reconstruct a phylogeny with Boletus reticuloceps (EU231968) as outgroup. The analysis revealed

127 five major clades corresponding with the taxonomic sections. Russula delica clustered with similar taxa in a same clade forming a sister clade with R. brevipes Peck with strong boot strap support.

Comments

Russula delica was first described from Sweden in 1838. It is widespread in the northern temperate zones, including Europe and Asia (Phillips, 2006). It has also been found from Pakistan in 1992 (Ahmad et al., 1997). It is commonly known as milk white brittle gill because of its white context (Nilson & Persson, 1977). It is mostly white with some ochoraceous tinge and a short robust stem. The species has undergone many taxonomic changes because of its close resemblance to other taxa. It can be confused easily with R. chloroides (Krombh.) Bres., R. pallidospora J. Blum ex Romagn. and R. flavispora Romagn. because of minor differences in their morphology (Haas, 1969).

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A B

C D

E Figure 63. Morphology of Russula delica. A–E. Basidiomata. A & B. LAH35067; C. LAH35068; D. LAH35069; E. LAH35065. Bars: A–E = 1.3 cm.

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Figure 64. Anatomy of Russula delica. A–F. LAH35068. A. Basidiospores; B. Basidia; Pleurocystidia; D. Cheilocystidia; E. Pileipellis; F. Stipitipellis. Bars: A–F = 10 µm.

130

KJ735006_Russula_sp KJ735007_Russula_sp 64 KJ735005_Russula_sp FJ845429_Russula_brevipes 87 KJ735017_Russula_sp KJ735014_Russula_sp JX630807_Russula_brevipes 89 DQ422005_Russula_aff_delica 51 72 HQ445587_Russula_cf_delica 71 JF834359_Russula_aff_brevipes 97 KF007188_Russula_cf_brevipes AY061671_Russula_delica EU819422Russula_brevipes_var_acrior 78 60 99 JX030213_Russula_aff_brevipes Lactarioides 94 Russula delica SJ51 70 KF432955_Russula_delica AF418605_Russula_delica KF007186_Russula_cf_brevipes 68 90 JF834356_Russula_aff_brevipes KF386757_Russula_brevipes 97 AF349714_Russula_brevipes 100 KF007187_Russula_cf_brevipes AF418604_Russula_chloroides KF432954_Russula_chloroides 98 69 JF834332_Russula_aff_chloroides 61 DQ422016_Russula_cf_chloroides 93 DQ658888_Russula_chloroides 88 EU819426_Russula_mariae EU819437_Russula_virescens Heterophyllae 85 AF418610_Russula_vesca 99 AF418609_Russula_heterophylla 100 GU371295_Russula_livescens AY061700_Russula_insignis Ingratae 99 AB211275_Russula_sororia 96 JN681168_Russula_cerolens 99 HQ604848_Russula_lutea 100 JF908682_Russula_amethystina Tenellae GU234063_Russula_chamiteae 99 FJ845432_Russula_decolorans EU597075_Russula_nigricans JF908669_Russula_adusta 99 Compactae EU526006_Russula_cascadensis 72 80 JF908673_Russula_anthracina EU231968_Boletus_reticuloceps Out group

0.05 Figure 65. Molecular phylogenetic analysis of Russula delica based on ITS sequences. The evolutionary history was inferred by the Maximum Likelihood method using Jukes-Cantor model. The analysis involved 44 nucleotide sequences. There were a total of 734 positions in the final dataset. Sequences generated from LAH35068 are marked with .

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Russula livescens (Batsch) Bataille, Fl. Monogr. Astérosporales: 76 (1908)

Jabeen et al., 2015 (Annexture)

Russula pakistanica nom. prov. (Figures 66–68)

Pileus 1–4.5 cm broad, depressed from the middle; dark brown (5YR1/2) from the centre with light brown (5YR5/4) to grayish brown (5YR5/2) tubeculate striations radially emerging out from the patches towards the margins on a reflective white context giving fibrillose appearance, surface dry; margins smooth, incurved when young, uplifted as infundibuliform at maturity. Lamellae up to 0. 25 cm broad, creamy white (2.5Y9/2), regular, adnexed, sub distant to close, edges entire. Lamellulae absent. Stipe 2.5–3.5 cm × 0.7–1 cm, central, cylindrical, slightly narrower towards the base; surface smooth to fibrillose, white to cream (2.5Y9/2) towards the apex and grayish brown (5YR5/2) towards the base.

Basidiospores [60/6/6] (5.6) 6.1–9.2 (9.4) × (5.4) 5.7–6.6 (7.5) µm, Q = (1) 1.03– 1.28 (1.56), avQ = 1.18 ± 0.17; globose to subglobose, ornamented, high warts with bifurcated apices are common accompanied with some low warts having rounded apices sometimes forming at the most a partial reticulum on the exine; warts up to 0.3 µm in trypan blue, apiculus prominent, guttulate, pale yellow in 5% KOH, amyloid in Malzer‟s reagent. Basidia (29) 29.7–38.9 (40.1) × (9.2) 9.4–11.3 (11.8) µm, clavate, thin walled, wall up to 3 µm thick, pale yellow in 5% KOH, guttulate. Pleurocystidia (64.7) 66.5–78.3 (80.7) × (7.5) 7.8–8.3 (9) µm, variable in shape, from fusiform, mucronate to rostrate, guttulate, pale yellow in 5% KOH. Cheilocystidia (49.7) 52.6–57.3 (59.9) × (8.9) 9.9– 10.85 (11.5) µm, fusiform, mucronate to rostrate, guttulate, pale yellow in 5% KOH. Pileipellis hyphae (2.1) 3.3–4.2 (4.5) µm wide, filamentous, branched, bifurcate terminals, pale yellow to transparent in 5% KOH, septa infrequent, clamp connections and pileocystidia not observed. Stipitipellis hyphal wall undulating, irregular in size, shape and diameter, (1.4) 1.6–7 (8) µm wide, cellular to filamentous, branched, bifurcate terminals, (1.4) 1.6–7 (8) µm wide, pale yellow to transparent, rarely septate, clamp connections not observed.

Material examined: Pakistan, Khyber Pakhtunkhwa province, Malakand division; Swat district; Kalam, 2200 m asl, on soil under Cedrus deodara, 3 Sep 2013, Sana Jabeen K8; SJ54 (LAH35005); 4 Sep 2013, Sana Jabeen K2-23; SJ74 (LAH35006); Mushkun, 2500

132 m asl, on soil under C. deodara, 21 Sep 2013 Sana Jabeen K4-26; SJ48 (LAH35007); Rawalpindi division & district, Kuzah Gali, 2500 m asl, on soil under C.s deodara, 21 Sep 2013, Sana Jabeen KG-3; SJ9 (LAH35004).

Morphological characterization of ectomycorrhiza (Figure 69)

Ectomycorrhizal system dichotomously ramified to coralloid with wooly appearance; up to 4 mm long; tips 0.5–2 mm long and up to 0.25 mm wide, apex 0.2 mm wide, base up to 0.25 mm wide; main axis 1 mm in diameter; light brown (10YR4/10) to dark brown (10YR3/8) or black at maturity; un ramified ends straight, cylindrical; Mantle silverish white to pale yellow (10YR8/8), distinct, reflective; host tissue visible under the mantle surface. Rhizomorphs frequent, smooth, pale yellow (5YR7/10) to transparent. Emanating hyphae abundant branched, pale yellow (5YR7/10) to transparent.

Mantle psedoparenchymatous in all layers, non-gelatinous, cells variable in shape.

Inner mantle layer pseudoparenchymatous, cells angular to rounded, elongated or lobed, tend towards the plectenchymatous state (mantle type L, Agerer, 1987–2002; Agerer & Rambold, 1998), cells membranaceously and plasmatically pale yellow to transparent,

10.7 ± 3.2 × 5 ± 0.6 µm, cell contents clear. Outer mantle layer pseudoparenchymatous, cells angular to rounded, elongated or lobed, tend towards the plectenchymatous state (mantle type L, Agerer, 1987–2002; Agerer & Rambold, 1998), cells membranaceously and plasmatically pale yellow to transparent, 10.6 ± 4.1 × 5 ± 0.4 µm, matrix clear.

Emanating hyphae straight, elongated, cylindrical, septate, clamped septa frequent, 2.2 µm in diameter, surface smooth, contents clear, ramification rather frequent, Y-shaped; hyphal endings simple to bifurcate, anastomosing not observed. Cystidia frequent, bottle shaped, 14.7 µm in length with strongly inflated base, upto 5.36 µm wide, a long straight neck 2.3 µm in diameter. Rhizomorphs frequent, hyphae branched, uniform and loosely arranged, intermingled, 2.7 µm in diameter, septate, clamp connections frequent, pale yellow to transparent.

Material examined: Pakistan, Khyber Pakhtunkhwa province, Hazara division; abbotabad district; Khanspur, 2400 m asl, asociated with Cedrus deodara, 19 May 2014 Sana Jabeen & Abdul Nasir Khalid Hp-14-3c; SA178 (LAH-EM5-2013); Malakand division; Swat district; Mushkun, 2500 m asl, associated with C. deodara, 21 Sep 2013 Sana Jabeen K4-100a (LAH-EM48-2013); Rawalpindi division & district, Patriata, 2200

133 m asl, associated with C. deodara, 21 Sep 2013, Sana Jabeen & Hassan Ayub Pt-3c; SA171 (LAH-EM3-2013); Pt; SA110 (LAH-EM49-2013); Pt-4; SA158 (LAH-EM50- 2013); Pt-7b; (LAH-EM51-2013).

Molecular phylogenetic characterization (Figures 70 & 71)

Consensus sequence of 611 base pairs was obtained by trimming the conserved motifs. BLAST of search revealed that the sequences generated from Russula pakistanica fruit body and ectomycorrhizal tissue were 95% identical to R. pseudopectinatoides (KM269079, KM269076 & KM269077) from China and 94% identical to R. pectinatoides Peck from USA (KF245518, EU819493 & KF245514) and Italy (JF908639) with 100% query cover and 0.0 E value. It also showed 94% identity to R. amoenolens Romagn. (GQ166870) from USA.

Sequence from LSU showed 97% identity to R. pectinatoides (KT933836) from USA and Sweden (DQ422026). It also showed 97% identity to R. granulata (KT933832) from USA with 94% query cover and 0.0 E value.

To reconstruct phylogeny, closely related ITS and LSU sequences were retrieved from the GenBank. Boletus reticuloceps (Zang et al.) Wang & Yao was chosen as out group in both, ITS and LSU sequence analysis (Shimono et al., 2004). were chosen as out group. Sequences of Russula pakistanica from Pakistan clustered in sect. Ingratae forming a sister clade with R. pectinatoides and allies with strong boot strap support.

Comments

Russula pakistanica is characterized by dark brown to grayish brown, weakly tuberculate pileus, creamy white adnexed lamellae, absence of lamellulae, globose to subglobose amyloid basidiospores with bifurcated warts on the exine, presence of hymenial cystidia, and filamentous pileipellis with bifurcated ends.

Russula pakistanica is distinct from R. pectinatoides Peck by its dark brown disc, dry pileus and pileal hyphae with bifurcate endings. Moreover, stipe surface layer of R. pectinatoides consists of longitudinal interwoven hyphae with regular elongated shape, while the hyphae of the stipe surface layer of R. pakistanica are highly variable in size, shape and diameter. Hyphal walls are not undulating. Hyphal endings are mostly branched and tapered (Adamčík et al., 2013).

134

Russula praetervisa Sarnari, another phylogenetically close relative of R. pakistanica resembles morphologically with R. pakistanica in having tuberculate striations on the pileus margin. It differs in the presence of reddish brown to blood red patches on the base of the stipe. Spore exine bears warts along with connectives (Sarnari, 1998; Das et al., 2013). The species also differs from R. pulverulenta and R. granulata by its non scurfy and non flocculose pileal surface (Shaffer, 1972).

Among the Asian species, R. dubdiana K. Das et al. (2013), a species from broad leaved forests in India can be separated from the present species by its viscid to glutinous pileus, smaller basidiospores (6–7 × 4–5.5 µm) with isolated cylindrical warts and a pileipellis with capitate pileocystidia. Russulla natarajanii K. Das et al. also originally described from Indian broad leaved forests is characterized by its white to yellowish white pileus (Das et al., 2006).

Russula fuscogrisea Petch, originally described from broad leaved forests of Sri Lanka differs in having isolated warts on basidiospore wall and shorter plurocystidia (50– 70 × 4–7.5 µm). Ixocutis with hyphae embedded in a thick, gelatinous matrix is another strong differentiating feature of R. fuscogrisea (Pegler, 1986). The other comparable species R. periglypta Berk. & Broome, from the same habitat in Sri Lanka and also known from southern India, has spinose spore wall and viscid, off white pileus with green tints (Pegler, 1986; Manimohan & Latha, 2011).

Russula punctipes Singer, from Yunnan, China has a stipe with pale blackish brown dots (Li, 2014). Russula pseudopectinatoides Li & Wen, a newly described species from China is distinct from R. pakistanica by its verrucose to conical warts on spore wall and pileipellis hyphae with obtuse to ventricose tips (Li et al., 2015)

Ectomycorrhizal features of Russula pakistanica are distinct from the ectomycorrhizae of related taxa. In R. amoenolins Romagn., cystidia on the mantle surface are flask shaped with an apical knob (Agerer, 1987–2006), while it is bottle shaped with a straight long neck having a blunt apex in R. pakistanica. The emanating hyphae have bifurcate terminal elements were also observed in the pileus epicutis of fruit body of R. pakistanica. Combination of all these characters of the basidiomata and EcM makes R. pakistanica distinct from other reported taxa. Molecular phylogenetic analysis also suggests it as a new taxon of the genus with a strong support.

135

All the specimen of R. pakistanica cited in this study have been collected from mixed coniferous forests of Pakistan. Its ectomycorrhiza has been found to be associated with Cedrus deodara in this study. Sequences (HG796931 & HG796943) have also been generated from ectomycorrhizal tissue of deciduous trees (Ilyas, 2013). Association of Russula pakistanica with conifers and deciduous trees suggests that the species has a broad host range.

136

Figure 66. Morphology of Russula pakistanica. A–E. Basidiomata. A & B. LAH35004 (holotype); C & D. LAH35005; E. LAH35006. Bars: A–E= 1.2 cm.

137

Figure 67. Scanning electron micrographs of basidiospores of Russula pakistanica . LAH35004.

138

Figure 68. Anatomy of Russula pakistanica. A–F. LAH35004. A. Basidiospores; B. Basidia; C. Pleurocystidia; D. Cheilocystidia; E. Stipitipellis; F. Pileipellis. Bar: A = 5 µm, B = 15 µm, C & D = 12 µm, E & F = 2.5 µm.

139

Figure 69. Ectomycorrhiza of Russula pakistanica. A–G. LAH-EM3-2013. A. Ectomycorrhizal system; B. Ectomycorrhizal root tip; C. Inner mantle layer; D. Outer mantle layer; E. Emanating hyphae; F. Cystidia. Bars: A = 0.5 µm, B = 2 µm, C & D = 11.5 µm; E = 13 µm; F = 0.8 µm; G = 11 µm.

140

KF245514 R pectinatoides EU819493 R pectinatoides 99 KJ834569 R pectinatoides KF245518 R pectinatoides GQ166870 R amoenolens 52 KJ834567 R pectinatoides KJ834571 R pectinatoides KM052566 R pectinatoides 36 JX425405 R pectinatoides 64 JX434670 R pectinatoides 46 39 JF273538 R pectinatoides KF318072 R pectinatoides 94 KF318063 R pectinatoides 75 DQ422026 R pectinatoides KF245531 R praetervisa UDB015983 R pectinatoides UDB011156 R pectinatoides 82 48 92 JF908639 R pectinatoides UDB019331 R praetervisa KJ834578 R praetervisa KF303600 R praetervisa KF303598 R praetervisa 35 KF303597 R praetervisa 100 KJ834614 R praetervisa KF303606 R praetervisa UDB019333 R praetervisa KF303599 R praetervisa 88 KM269079 R pseudopectinatoides KM269077 R pseudopectinatoides 99 87 KM269078 R pseudopectinatoides KU535608 R pakistanica R pakistanica SA171 HG796943 R pakistanica 100 KU535609 R pakistanica KT834641 R pakistanica SJ48 KT834639 R pakistanica SJ54 90 Ingratae KT834640 R pakistanica SJ74 KT834638 R pakistanica SJ9 HG796931 R pakistanica DQ422024 R illota 59 100 HQ677769 R illota KF245526 R illota 72 KF245509 R illota AF418614 R grata 32 KF245532 R grata 99 KF245527 R laurocerasi AY239335 Gymnomyces parksii 46 82 KC152107 Gymnomyces fallax 97 JX425383 R foetens KF245500 R cf. subfoetens LN714598 R foetens 69 AF418613 R foetens 99 AF418613 R foetens 72 AY061706 R pectinata 99 GU222264 R amoenolens 21 93 AF418615 R amoenolens KF245505 R cerolens 19 97 KF245524 R cerolens 89 JN681168 R cerolens AB211275 R sororia 87 JQ622327 R amoenolens 98 EU598186 R pulverulenta 50 AY061736 R pulverulenta 22 98 AY061700 R insignis KF850404 R insignis 93 GU371295 R livescens KJ834624 R granulata 39 KF359618 R granulata 22 EU598189 R granulata 100 EU598191 R granulata EU598188 R granulata JQ272365 R granulata 100 AF418610 R vesca AF418609 R heterophylla Heterophyllae 98 AF418611 R parazurea AF418612 R aeruginea Germany 73 99 KF245525 R cf aeruginea 76 GU234063 R chamiteae 81 FJ845432 R decolorans JQ888199 R paludosa DQ422022 R risigallina Tenellae 100 HQ604848 R lutea 94 JF908682 R amethystina 51 KF002762 R gilva AB291768 R adusta AB291765 R densifolia Compactae 99 AB291729 R nigricans 63 66 AB291751 R subnigricans EU231968 Boletus reticuloceps Outgroup

0.05

Figure 70. Molecular phylogenetic analysis of Russula pakistanica based on ITS sequences. The evolutionary history was inferred by the Maximum Likelihood method using General Time Reversible model. The analysis involved 89 nucleotide sequences. There were a total of 782 positions in the final dataset. Sequences generated during this investigation are marked with .

141

99 KT933826 Russula crustosa 77 KT933822 Russula crustosa 54 EU598167 Russula sp 33 KT933866 Russula mustelina 25 DQ422014 Russula virescens DQ422007 Russula parazurea 35 22 DQ422012 Russula ochrospora 95 KT933834 Russula aff azurea KT933870 Russula cf atroglauca 27 90 JX391970 Russula sp 75 KT933867 Russula grisea 79 JN389003 Russula columbicolor DQ422006 Russula heterophylla 40 KT933839 Russula vesca 80 57 AF218548 Russula brunneola 57 DQ422018 Russula vesca 31 AF2878881 Russula compacta

97 EU598164 Russula nitida 92 KT933811 Russula cf firmula AF218547 Russula radicans 31 KT933808 Russula redolens 29 97 KT933825 Russula redolens KT933832 Russula granulata 73 DQ422026 Russula pectinatoides 62 KT933836 Russula pectinatoides 50 Russula pakistanica SJ9 33 JN389006 Russula tsokae KT933849 Russula laurocerasi 33 27 DQ422024 Russula illota 30 KT933877 Russula foetens 22 AF218546 Russula foetentula KT933813 Russula amoenolens 80 DQ422023 Russula cf foetens AF265542 Martellia fallax 82 KP859294 Gymnomyces nondistincta DQ422029 Russula albonigra 95 DQ421998 Russula acrifolia AY293209 Russula exalbicans KT933871 Russula cuprea 99 28 KT933855 Russula velutipes 56 KT933817 Russula rubellipes DQ421984 Russula ochricompacta 98 DQ421986 Russula ochricompacta KF112454 Boletus reticuloceps

0.02 Figure 71. Molecular phylogenetic analysis of Russula pakistanica based on LSU sequences. The evolutionary history was inferred by the Maximum Likelihood method using General Time Reversible model. The analysis involved 44 nucleotide sequences. There were a total of 915 positions in the final dataset. Sequences generated from LAH35004 are marked with .

142

Russula rubecola nom. prov. (Figures 72 & 73)

Pileus up to 8.2 cm broad, depressed from the middle, pileipellis pale yellow (5Y8/6) from the centre, bright red (7.5R5/16) towards the margins, becoming blackish red (2.5YR2/8) with time, surface dry, smooth becoming striated towards the margin, margins incurved when young becoming straight at maturity. Lamellae up to 0.3 cm broad, white to pale yellow (5Y8/6), regular, adnexed, subdistant to close, edges entire. Lamellulae absent. Stipe 6.2 × 1.5 cm, white, central, cylindrical, slightly narrower towards the apex. Surface smooth to finely wrinkled or fibrillose.

Basidiospores [40/2/2] (10.5) 11.3–11.5 (12.3) × (7.8) 8.4–9.4 (9.8) µm, Q = (1.10) 1.20–1.32 (1.58), avQ = 1.24; globose to subglobose, echinulate; apiculus prominent, pale yellow to hyaline in 5% KOH, amyloid in Melzer‟s reagent. Basidia (38.3) 41.4–43.2 (44.6) × (10.1) 12.5–14.7 (16.1) µm, clavate, wall up to 3 µm, hyaline in 5% KOH. Pleurocystidia (89.4) 91.4–106.7 (110.2) × (10.2) 11.6–13.2 (14.7) µm, fusoid, pale yellow to hyaline in 5% KOH. Cheilocystidia (44.0) 54.6–90.2 (131.3) × (10.2) 13.4–18.3 (24.1) µm, fusiform, guttulate, pale yellow to hyaline in 5% KOH. Pileipellis hyphae (2.0) 2.1–2.4 (2.6) µm wide, filamentous, branched, hyaline in 5% KOH, septa frequent, clamp connections common. Stipitipellis hyphae (1.3) 1.6–2.4 (3.2) µm wide, filamentous, branched, hyaline, clamped septa common.

Material examined: Pakistan, Khyber Pakhtunkhwa province, Malakand division, Swat district, Mashkun 2500 m asl, on soil under Cedrus deodara, Aug 2014, Sana Jabeen 5; SJ106 (LAH35070); Abbotabad district, Shimla Hill, 1500 m asl, on soil under C. deodara, 6 Aug 2014, Sana Jabeen & Abdul Nasir Khalid SJ105 (LAH35071).

Molecular phylogenetic characterization (Figure 74)

Sequencing of the Russula rubecola PCR products using ITS1F and ITS4 primers yielded 512–739 base pairs. A consensus sequence of 638 base pairs was obtained by trimming the motifs. BLAST revealed it as 89% identical to Russula sp. sequences (KR082881 & KR082881) from China. It also showed 88% similarity with Russula sp. (KM576560) from Hungry with 97% query cover and 0.0 E value.

Published sequences from different Russula sections were included to reconstruct a phylogeny with Boletus reticuloceps (Zang et al.) Wang & Yao (EU231968) as out group. The analysis revealed four major clades corresponding with the taxonomic

143 sections. Species within these clades were clustered with strong bootstrap, supported by morphological characters. Russula rubecola clustered with sequences from China and Hungary in a same clade. It gets separated forming its own lineage within section Tenellae with a strong boot strap value.

Comments

Russula rubecola is characterized by its smooth, depressed, yellowish pileus from the middle, bright red and striated towards the margins becoming blackish red with age. It is also characterized by its yellowish hymenium. The species is closely related to R. maculata Quél. Morphologically R. maculata is distinct from R. rubecola by its pink to reddish pileus, comparatively small and wide stipe with pink tint, sometimes rusty spotted or browning, spore size also comparatively smaller than that of R. rubecola, and no septa in the pileal hyphae (http://www.rogersmushrooms.com/). It is also closely related to R. chiui Li & Wen, a newly reported taxon from India. Russula chiui is distinct from R. rubecola by its non striate pink to peach red or yellow orange pileus, comparatively smaller spores (8.5–10.5 × 7–8.5) µm and absence of clamp connections from all types of hyphae (Li et al., 2015). Molecular analysis based on ITS sequences also proves it distinct from all other taxa within Tenellae.

144

\

A

B

Figure 72. Morphology of Russula rubecola. A & B. Basidiomata LAH35071 (holotype). Bars: A & B = 1.2 cm.

145

Figure 73. Anatomy of Russula rubecola. A–F. LAH35071. A. basidiospores; B. cystidia; C. basidia; D. pileipellis; E. stipitipellis. Bars: A–E = 10 µm.

146

JQ888203_Russula_vinosa AF418638_Russula_vinosa AY061724_Russula_vinosa 100 JX029957_Russula_sp JF908703_Russula_vinosa JF834328_Russula_aff_vinosa 90 FJ845436_Russula_occidentalis 58 AY534206_Russula_occidentalis AF418630_Russula_veternosa KC797152_21548_Russula_firmula 36 92 JF834342_Russula_aff_firmula 64 KJ867372_Russula_firmula 88 AF418631_Russula_firmula 80 96 AY061653_Russula_amethystina 66 AY245542_Russula_californiensis GU234063_Russula_chamiteae 46 90 FJ845432_Russula_decolorans KF850410_Russula_postiana KF002762_Russula_gilva 98 78 KF679818_Russula_lutea 100 AF230898_Russula_postiana 74 72 UDB000897_R_postiana 50 HQ604848_Russula_lutea DQ422022_Russula_risigallina Tenellae 26 AY061713_Russula_risigallina 100 78 JF908702_Russula_velenovskyi JF908706_Russula_puellula KM576540_Russula_sp 100 100 KC952701_Fagus_sylvatica KC952700_Fagus_sylvatica 46 KC952676_Fagus_sylvatica AY061710_Russula_puellula 94 DQ422015_Russula_cf_maculata 100 AY061688_Russula_maculata JN129407_Russula_sp 76 KR082870_Russula_maculata 100 KR082881_Russula_sp Russula rubecola SJ105 Russula rubecola SJ106 100 84 KF810136_Russula_subrubescens 62 EU569276_Russula_sp KM576560_Russula_sp 92 AF495464_Russula_sp AF495465_Russula_sp 64 JF908710_Russula_dryadicola KF225495_Russula_sp KF225492_Russula_sp 46 76 KF225494_Russula_sp 100 JF908673_Russula_anthracina 74 JF834363_Russula_acrifolia 58 EU303008_Russula_dissimulans 76 JQ888194_Russula_adusta Compactae 100 JF834355_Russula_albonigra KC581314_Russula_nigricans 58 AB291765_Russula_densifolia 100 AF418610_Russula_vesca 46 AF418609_Russula_heterophylla 44 EU819426_Russula_mariae 76 Heterophyllae 76 EU819437_Russula_virescens AF418612_Russula_aeruginea 60 AF418611_Russula_parazurea 60 JQ622344_Russula_pulverulenta 70 100 AY061700_Russula_insignis GU371295_Russula_livescens 100 KF002757_Russula_subfoetens Ingratae KF245530_Russula_foetens 76 HQ677769_Russula_illota 50 56 KF245527_Russula_laurocerasi EU231968 Boletus reticuloceps Out group

0.05 Figure 74. Molecular phylogenetic analysis of Russula rubecola based on ITS sequences. The evolutionary history was inferred by the Maximum Likelihood method using General Time Reversible model. The analysis involved 69 nucleotide sequences. There were a total of 842 positions in the final dataset. Sequences generated during this study are marked with .

147

Genus Suillus

Suillus convexatus nom. prov. (Figures 75 & 76)

Etymology: The specific epithet refers to the convex pileus.

Pileus up to 4.2 cm, convex, spherical to irregular, surface dry, rough, off white (5Y9/2) to light brown (10YR8/4) becoming dark brown (7.5YR5/6) towards age with some grayish (10YR5/2) patches. Hymenium yellowish brown (2.5YR5/10) to blackish, pores frequent, up to 0.5 cm deep. Stipe 2 × 0.7 cm, base slightly narrow up to 0.4 cm towards the base, light brown (10YR8/6), rough and dry.

Basidispores [30/2/1] (9.5) 9.8–11.4 (12.1) × (3.8) 3.9–5.2 (5.3) µm, Q = (2.06) 2.09–2.20 (2.90), avQ = 2.08, ellipsoid, smooth, epiculus prominent. Basidia 40.3–44.7 × 5.7–6.5 µm, clavate, thin walled, guttulate, contents visible. Cystidia 39.6–45.7 × 4.9–5.7 µm, cylindrical to subclavate, thin walled, hyaline in 5% KOH. Pileipellis hyphae (5.9) 7.2–8.7 (10.1) µm, filamentous, clavate terminals, hyaline in 5% KOH. Stipitipellis hyphae (4.9) 5.7–6.9 (9.0) µm, filamentous, clavate terminals, hyaline in 5% KOH.

Material examined: Pakistan, Khyber Pakhtunkhwa province, Malakand division, Swat district, Kalam, 2400 m asl, on soil under Cedrus deodara 3 Sep 2013, Sana Jabeen K2- 17; SJ62 (LAH35072).

Molecular phylogenetic characterization (Figure 77)

Sequencing of the PCR products of ITS region of Suillus convexatus yielded 727– 743 base pairs by using ITS1F and ITS4 primers. Consensus sequence of 628 base pairs was obtained by trimming the motif and BLAST searched at NCBI. It showed 97% identity to Suillus indicus B. Verma & M.S. Reddy (as Suillus sp.) (KJ675500– KJ675502) from India, and 92% identity to Suillus quiescens T.D. Bruns & Vellinga (NR_119750, GQ249390, JN858077 & GQ249393) from USA with 99% query cover and 0.0 E value.

To reconstruct phylogeny, closely related ITS sequences were retrieved from the GenBank. Rhizopogon luteolus Fr. (JQ888192) was chosen as out group. The sequence generated during this study gets separated from Indian and USA taxa forming its own lineage in the same clade with a strong boot strap value.

148

Comments

Suillus convexatus is characterized by its convex off white to brownish pileus, a short stipe becoming narrower towards the base, smooth ellipsoid spores and cylindrical hymenal cystidia. It can be compared with S. indicus from India. Suillus convexatus differs from Indian specimen on the basis of its size and color of the pileal surface. Suillus indicus bears comparatively short basidiomata with a very small and narrow stipe, a reddish brown pileal surface and yellow hymenium. Presence of veil remnants and upturned margins also make it distinct from S. convexatus (Verma & Reddy, 2015). Suillus convevxatus also differs from S. marginielevatus S. Sarwar, Khalid & Dentinger, a newly reported taxon from Pakistan on the basis of its margins which are upturned and the large size of basidiomata as well as yellow hymenium (Sarwar et al., 2015). Molecular data analysis based on ITS region also support S. convexatus as a distinct taxon.

149

Figure 75. Morphology of Suillus convexatus. Basidiomata LAH35072 (holotype). Bar = 0.6 cm.

150

Figure 76. Anatomy of Suillus convexatus. A–E. LAH35072. A. Basidiospores; B. Cystidia; C. Basidia; D. Pileipellis; E. Stipitipellis. Bars: A–E = 10 µm.

151

69 KM882911_Suillus_granulatus 88 KM882910_Suillus_granulatus 74 KM882912_Suillus_granulatus L54121_Suillus_cf_granulatus 81 HM347658_Suillus_collinitus AY953421_Suillus_collinitus 68 81 HM347660_Suillus_collinitus_var_velatipes KM882908_Suillus_triacicularis KM882907_Suillus_triacicularis 87 99 KF977189_Suillus_sp_ KF977188_Suillus_sp JN858077_Suillus_quiescens GQ249390_Suillus_quiescens 94 GQ249393_Suillus_quiescens_ 96 GQ249401_Suillus_quiescens NR_119750_Suillus_quiescens L54094_Suillus_pungens 99 JQ310819_Suillus_pungens Suillus convexatus SJ62 55 KJ675502_Suillus_sp 70 KJ675501_Suillus_sp KJ675500_Suillus_sp 96 KJ361512_Suillus_sp KP792121_Suillus_sp 67 KP792120_Suillus_sp 78 KJ361513_Suillus_sp L54118_Suillus_cf_placidus AB284443_Suillus_placidus 99 KJ415101_Suillus_placidus KM882921_Suillus_placidus L54092_Suillus_cothurnatus AJ419218_Suillus_cothurnatus 97 L54088_Suillus_subluteus 70 KP698003_Suillus_subluteus AM502982_Suillus_sp KJ472765_Suillus_sp KM882918_Suillus_sibiricus KF699850_Suillus_sp JN119753_Suillus_sibiricus KM882916_Suillus_sibiricus 72 JN119752_Suillus_sibiricus KM248957_Suillus_americanus KT336209_Suillus_cf._sibiricus L54103_Suillus_americanus JQ711962_Suillus_flavidus JQ711871_Suillus_flavidus JQ711869_Suillus_flavidus 73 JQ711908_Suillus_flavidus JQ711885_Suillus_flavidus L54115_Suillus_umbonatus JQ888192_Rhizopogon_luteolus

0.02 Figure 77. Molecular phylogenetic analysis of Suillus convexatus based on ITS sequences. The evolutionary history was inferred by the Maximum Likelihood method using General Time Reversible model. The analysis involved 51 nucleotide sequences. There were a total of 423 positions in the final dataset. Sequences generated during present investigation are marked with .

152

Genus Tricholoma

Tricholoma terreum (Schaeff.) P. Kumm., Führ. Pilzk. (Zerbst): 134 (1871)

(Figures 78 & 79)

Pileus up to 5 cm, convex to flat, becoming infundibuliform when mature, surface dry, dark brown (2.5YR1/2) becoming light brown (2.5YR4/8) towards the margins. Lamellae off white, distant, up to 0.6 cm broad, adnexed, edges entire, margins wavy. Lamellulae variable in size. Stipe 6 × 1.4 cm, off white with some grayish (5Y6/2) patches.

Basidiospores [30/1/1] (6.1) 6.6–7.5 (8.0) × (4.9) 5.2–5.7 (6.1) µm, Q = (0.25) 0.26–0.28 (0.36), avQ = 0.28, globose to sub globose, smooth, epiculus prominent, hyalone in 5% KOH. Basidia (10.2) 11.7–13.4 (14.0) × (5.2) 5.7–6.5 (7.1) µm, clavate, thin walled, contents visible, hyaline in 5% KOH. Hymenal cystidia (9.9) 10.4–12.0 (12.7) × (4.9) 5.4–6.3 (6.8) µm, cylindrical to subclavate, thin walled, hyaline in 5% KOH. Pileipellis hyphae (3.9) 4.2–8.7 (10.1) µm, filamentous, frequently septate, hyaline in 5% KOH. Stipitipellis hyphae (2.8) 4.0–7.3 (8.1) µm, filamentous, clavate to fusiform terminals, hyaline in 5% KOH.

Material examined: Pakistan, Punjab province, Rawalpindi division & district, Kuza Gali, 2500 m asl, on soil under Cedrus deodara, 21 Sep 2013, Sana Jabeen KG-4b; SJ45 (LAH35073).

Molecular phylogenetic characterization (Figure 80)

Sequencing of the PCR products of ITS region of Tricholoma terreum yielded 728–807 base pairs by using ITS1F and ITS4 primers. Consensus sequence of 531 base pairs was obtained by trimming the motif and BLAST searched at NCBI. It showed 99% identity to T. terreum from China (EU439336), Denmark (EU653299 & EU653300), Germany (LT000098) and Sweden (LT000193) with 100% query cover and 0.0 E value.

To reconstruct phylogeny, closely related ITS sequences were retrieved from the GenBank. Lyophyllum caerulescens Clémençon (KP192628) was chosen as out group. The sequence generated during this study clustered with similar taxa in same clade with strong boot strap support.

153

Comments

Tricholoma terrum was first described as Agaricus terreus in 1762. Its current name was given by German Paul Kummer in 1871. It is commonly known as "the gray knight or dirty tricholoma" due to its discolored gills (Lamaison & Polese, 2005; Phillips, 2006). It is mostly reported from Europe but records have also been found from other parts of the world (Grey, 2005; Keane et al., 2000; Bessette et al., 2013). It has also been recorded from Pakistan (Ahmad et al., 1997). The species is regarded as a good edible, but recent chemical tests show that the species may contain toxins, so its edibility is not recommended (Zeitlmayr, 1976).

154

Figure 78. Morphology of Tricholoma terreum. Basidiomata LAH35073. Bar = 0.6 cm.

155

Figure 79. Anatomy of Tricholoma terreum. A–F. LAH35073. A. Basidiospores; B. Basidia; C & D. Hymenal cystidia; E. Stipitipellis; F. Pileipellis. Bars: A = 4 µm; B–D = 2.5 µm; E & F = 7.5 µm.

156

EU439321 Tricholoma terreum KM085374 Tricholoma terreum EU439322 Tricholoma terreum EU439336 Tricholoma terreum 974573944 Tricholoma terreum EU653299 Tricholoma terreum LT000098 Tricholoma terreum EU439326 Tricholoma terreum EU439330 Tricholoma terreum EU439318 Tricholoma terreum EU439331 1531 Tricholoma terreum EU439327 Tricholoma terreum LT000193 Tricholoma terreum Tricholoma terreum SJ45 EU653300 Tricholoma terreum EU439341 Tricholoma terreum EU439328 Tricholoma terreum 100 EU439334 Tricholoma terreum JN389311 Tricholoma terreum JN389309 Tricholoma terreum JN389307 Tricholoma terreum AF062614 Tricholoma terreum GU060265 Tricholoma inocybeoides 62 99 GU060267 Tricholoma inocybeoides GU060266 1525 Tricholoma inocybe KJ705244 Tricholoma cingulatum HQ184100 Tricholoma cingulatum GU060262 Tricholoma cingulatum 99 KU695460 Tricholoma cingulatum 98 AF377198 Tricholoma cingulatum AF349697 Tricholoma cingulatum GU060263 Tricholoma cingulatum GU060275 Tricholoma argyraceum GU060277 Tricholoma argyraceum GU060271 Tricholoma argyraceum GU060270 Tricholoma argyraceum GU060269 Tricholoma argyraceum GU060278 Tricholoma argyraceum GU060273 Tricholoma argyraceum GU060276 Tricholoma argyraceum 63 GU060272 Tricholoma argyraceum GU060274 Tricholoma argyraceum 63 GU060268 Tricholoma argyraceum KT800072 Tricholoma cf myomyces 84 KT800281 KT800166 Tricholoma cf myomyces KT800259 Tricholoma cf myomyces JN389291 Tricholoma myomyces AF377210 Tricholoma myomyces 83 KP454018 Tricholoma myomyces JN389299 Tricholoma myomyces JN389294 Tricholoma myomyces 61 FJ845443 Tricholoma myomyces KP192628 Lyophyllum caerulescens Out group

0.01 Figure 80. Molecular phylogenetic analysis of Tricholoma terreum based on ITS sequences. The evolutionary history was inferred by the Maximum Likelihood method using Jukes-Cantor model. The analysis involved 54 nucleotide sequences. There were a total of 543 positions in the final dataset. Sequences generated during present investigation are marked with .

157

Genus Gyromitra

Gyromitra khanspurensis nom. prov. (Figures 81 & 82)

Etymology: The specific epithet refers to the site of collection.

Ascomata upto 3.4 cm high, Hymenium free from the hymenophore; irregular, up to 2 cm high; 3.5 cm wide at the widest point; yellowish brown (5YR7/12) to brown (2.5YR5/10); surface smooth to wrinkled, becoming more wrinkled with age. Hymenophore 3.5 × 1 cm; base slightly wider up to 1.3 cm, off white, surface smooth.

Ascospores [40/2/1] (14.2) 15.0–16.6 (17.3) × (7.1) 8.4–8.5 (8.7) µm, Q = (1.76) 1.8-1.9 (2), avQ = 1.8; smooth, ellipsoid, guttulate, short apiculus. Asci long up to 219 µm, wide up to 9.9 µm wide; 8 spored; cylindrical having a long narrow base; hyaline in 5% KOH. Paraphyses long upto 141µm; wide upto 6.7 µm, clavate becoming narrow towards the base; hyaline in 5% KOH.

Material examined: Pakistan, Khyber Pakhtunkhwa province, Abbottabad district, Khanspur, 2575 m asl, associated with Cedrus deodara (Roxb. ex D. Don) G. Don), 19 May 2014, Lubna Choudry and Abdul Nasir Khalid SJ95; MP2 (LAH35074).

Molecular phylogenetic characterization (Figure 83)

Sequencing of the PCR products of ITS region of Gyromitra khanspurensis yielded 757–758 base pairs by using ITS1F and ITS4 primers. A consensus sequence of 758 base pairs was BLAST searched at NCBI. It showed 89% identity to G. gigas (Krombh.) Cooke (JF908781) from Italy, 85–87% to G. slonevskii V.P. Heluta (JQ691488- JQ691490) from Ukraine and 99% to G. esculenta (Pers.) Fr. (KM204670– KM204673) from USA.

To reconstruct phylogeny, closely related ITS sequences were retrieved from the GenBank. Taxa from Morchella Dill. ex Pers. (KM252950 and KP670934) were chosen as out group. The sequences from Pakistan clustered in a same clade with G. gigas and G. slonevskii but it got separated from both these taxa forming a sister lineage of its own with G. gigas with a strong (100%) boot strap support.

158

Comments

Gyromitra khanspurensis nom. prov. is characterized by its yellowish brown wrinkled hymenium free from hymenophore, ellipsoid, hyaline and long clavate asci. Morphologically it is closely related to G. gigas, but differs by its size of ascospores which are comparatively larger (26–40 × 11.5–15 µm) in G. gigas (Kuo, 2012). Morphologically, G. khanspurensis is also distinct from G. slonevskii. Gyromitra slonevskii bears a brown to liver colored cap and white to brownish, rough and ribbed, slightly compressed stipe. size is also comparatively larger (27–35 × 11–15) um in G. slonevskii (Hetula, 2001). Molecular dataset comparisons also suggest G. khanspurensis as a distinct taxon with a strong boot strap value. Ecologically the species is saprobic but it is considered as mycorrhizal like true morals having both traits in the course of its life cycle (Harmaja, 1973).

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Figure. 81. Morphology of Gyromitra khanspurensis. Ascomata LAH35074 (holotype). Bar = 0.5 cm.

160

Figure 82. Anatomy of Gyromitra khanspurensis. A–C. LAH35010. A. Ascospores; B. Asci; C. Paraphyses. Bars: A–C = 10 µm.

161

KC152110 KC152109 Gyromitra infula 18 KJ146709 Gyromitra infula 60 GQ304944 Gyromitra infula 100 DQ384573 Gyromitra infula FJ859347 Gyromitra infula 100 GQ377484 Gyromitra infula 52 JF773588 Gyromitra infula EU837202 Gyromitra californica GQ304943 Gyromitra sphaerospora 100 26 80 EU837203 Gyromitra californica 94 EU837204 Gyromitra californica 100 EU784275 Hydnotrya michaelis EU784274 Hydnotrya michaelis 92 JF908765 Hydnotrya cerebriformis 88 GQ140236 Hydnotrya cerebriformis KC152119 Hydnotrya cerebriformis 42 100 KC152120 Hydnotrya cerebriformis 92 KC152118 Hydnotrya cerebriformis 56 GU373505 100 JN043313 Gyromitra esculenta KF692071 Gyromitra esculenta 100 JN043315 Gyromitra esculenta KM204672 Gyromitra esculenta 100 JX310422 Gyromitra sp KM204670 Gyromitra esculenta 98 KM204674 Gyromitra esculenta KM204673 Gyromitra esculenta KM204671 Gyromitra esculenta 100 Gyromitra khanspurensis SJ95 JF908781 Gyromitra gigas 82 JQ691489 Gyromitra slonevskii 100 JQ691488 Gyromitra slonevskii 100 JQ691490 Gyromitra slonevskii KM252950 Morchella fluvialis Out group 100 KP670934 Morchella crassipes

0.1

Figure 83. Molecular phylogenetic analysis of Gyromitra khanspurensis based on ITS sequences. The evolutionary history was inferred by the Maximum Likelihood method using Jukes-Cantor model. The analysis involved 36 nucleotide sequences. There were a total of 1337 positions in the final dataset. Sequences generated from LAH35074 are marked with .

162

Genus Morchella

Morchella pakistanica nom. prov. (Figures 84 & 85)

Etymology: The specific epithet refers to the name of the country.

Ascomata 4.5 cm high. Hymenium clustered on the top of the hymenophore up to 0.3 cm high and 2 cm broad, grayish black (7.5YR2/2). Hymenophore 4.2 cm high; 1.5 cm from centre, becoming narrower towards the base, base upto 1 cm wide, off white 5Y9/2 when young becoming brownish (5YR4/6) in the form of fine dots.

Ascospores [20/1/1] (7.3) 7.9–9.6 (9.8) × (4.4) 5.0–6.1 (6.4) µm, Q = (1.39) 1.46– 1.66 (1.86), avQ = 1.56; sub globose to ellipsoid; smooth; contents homogeneous, hyaline in 5% KOH. Asci upto 131 µm long and 8.1 µm wide at the widest point; 8 spored, cylindrical; hyaline in 5% KOH. Paraphyses upto 121 µm long and 5.7 µm wide, cylindrical; clavate, hyaline in 5% KOH.

Material examined: Pakistan, Khyber Pakhtunkhwa province, Abbottabad district, Khanspur, 2575 m asl, associated with Cedrus deodara (Roxb. ex D. Don) G. Don), 19 May 2014, Sana Jabeen SJ121; MP (LAH35075).

Molecular phylogenetic characterization (Figures 86)

Sequencing of the PCR products of ITS region of Morchella pakistanica yielded 833–835 base pairs by using ITS1F and ITS4 primers. A consensus sequence of 736 base pairs was obtained by trimming at conserved motifs and BLAST searched at NCBI. It showed 99% identity to M. cf. inamoena (KM588021) from Spain, M. elata Fr. (KM588019) from Switzeland, M. gigas (Batsch) Pers. from Germany (JQ691483, AJ543743 & AJ543742), India (GQ228475) and Kazakhstan (JQ691494), M. semilibera DC. from China (JQ723023), France (KJ174320 & KM587981) and Japan (LC028919) with 89–100% query cover and 0.0 E value.

To reconstruct phylogeny, closely related ITS sequences were retrieved from the GenBank. (O.F. Müll.) Sw. (AJ544206) was chosen as out group. The sequences from Pakistan diverged from other taxa forming their own lineage with 88% boot strap value.

163

Comments

Morchella pakistanica is characterized by its grayish hymenium at the top of the hymenophore, off-white stipe becoming brownish when dry and sub globose to elliptical smooth hyaline spores. It can be compared with a closely taxon; M. semilibera mostly foundin Europe, rare in Asia and not known to accur in America (Moreau et al., 2014). consists of a campanulate half free apothecial margins a long slender stipe (Moreau et al., 2014). Morchella pakistanica bears a small hymenium on a wide hymenophore becoming narrower towards the base in the form of a cone. Morchella semilibera has many synonyms as indicated by Moreau et al. (2014), and the ITS sequence from neotype of M. semilibera (KJ174320) clustered with other similar taxa from different countries. Morchella pakistanica diverged from all these taxa with a strong boot strap value suggests it as a distinct taxon.

As far as the trophic status of the Morchella Dill. ex Pers. is concerned, several reports suggest that in stable forest ecosystems, the life cycle of this genus includes both saprobic and mycorrhizal phases, oscillating in a biannual cycle (Buscot & Roux, 1987; Buscot, 1989; Buscot & Bernillon, 1991). Dahlstrom et al. (2000) tested the ability of Morchella to form mycorrhizae with various tree species in lab conditions and their results sugget that different members of Morchella successfully synthesized ectomycorrhizae with members of Pinaceae. Based on these findings, Morchella is treated as an ectomycrrhizal symbiont in this study.

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Figure. 84. Morphology of Morchella pakistanica. Ascoma LAH35075 (holotype). Bar = 0.3 cm.

165

Figure 85. Anatomy of Morchella pakistanica. A–C. LAH35075. A. Ascospores; B. Asci; C. Paraphysis. Bars: A–C= 10 µm.

166

KM588001 Morchella semilibera KM588007 Morchella semilibera JQ691483 Morchella gigas AJ543742 Morchella gigas KM588021 Morchella cf inamoena 62 JQ723024 Morchella semilibera LC028919 Morchella semilibera AJ543743 Morchella gigas KC464349 Morchella gigas 88 JQ691494 Morchella gigas KJ174320 Morchella semilibera KM588019 Morchella pakistanica SJ121 80 Morchella pakistanica SJ121b 100 KM204688 Morchella populiphila KM204689 Morchella populiphila 98 KM204690 Morchella populiphila KM587961 Morchella populiphila JQ723033 Morchella populiphila JQ723029 Morchella punctipes 46 JQ723025 Morchella punctipes JQ723026 Morchella punctipes JX069622 Morchella punctipes 96 JX069619 Morchella punctipes JX069617 Morchella punctipes JQ723028 Morchella punctipes 38 DQ257332 Morchella costata DQ257333 Morchella costata 100 DQ257334 Morchella costata EF080998 Morchella costata KM588006 KM588011 Morchella deliciosa JQ321881 Morchella capitata JQ618591 Morchella capitata 68 100 JQ618592 Morchella capitata 100 JQ618593 Morchella capitata JQ321880 Morchella capitata JQ723047 Morchella capitata 30 JQ723038 Morchella sextelata JQ321877 Morchella sextelata 88 50 KM587991 Morchella sextelata JQ723039 Morchella sextelata 78 JQ723036 Morchella sextelata JQ723034 Morchella sextelata 72 34 JQ723035 Morchella sextelata 22 AJ698475 AJ544206 Verpa conica Out group

0.05 Figure 86. Molecular phylogenetic analysis of Morchella pakistanica based on ITS sequences. The evolutionary history was inferred by the Maximum Likelihood method using Jukes-Cantor model. The analysis involved 47 nucleotide sequences. There were a total of 774 positions in the final dataset. Sequences generated from LAH35075 are marked with .

167

Genus Verpa

Verpa asiatica nom. prov. (Figures 87 & 88)

Etymology: The specific epithet refers to the geological region of occurrence.

Ascomata 4.5–5.5 cm high. Hymenium free from the hymenophore; 1.7–2.2 cm high; 1.5–1.9 cm wide at the widest point; light brown (2.5Y7/4) ridges and grayish (5YR4/2) furrows when young becoming gray (7.5YR3/2) to dark brown (7.5YR3/6) at maturity; obtusely cylindrical, pitted and ridged, with approximately 2–6 vertical ridges. Hymenophore 4.1–5.3 cm high; 1–1.2cm wide; base up to 1.5 cm wide; off white (7.5Y9/2) or yellowish (2.5Y8/10) to brown (10YR6/8); hollow; basally clavate to subclavate; surface finely granular.

Ascospores [60/4/2] (60.1) 67.1–74.3 (75.7) × (18.8) 20.1– 22.2 (22.7) µm; oblong; smooth; contents homogeneous. Asci 24–36 µm wide at the widest point; two spored, three spored also observed; cylindrical having a long narrow base; hyaline in 2% KOH. Paraphyses 5–7 µm; cylindrical; apices generally rounded or sub fusiform; septate; hyaline in 2% KOH.

Material examined: Pakistan, Khyber Pakhtunkhwa province, Abbottabad district, Khanspur, 2575 m asl, associated with Cedrus deodara (Roxb. ex D. Don) G. Don), 18 May 2014, Lubna Choudry and Abdul Nasir Khalid SJ120 (LAH35076); 19 May 2014, Abdul Nasir Khalid, Lubna Chaudry and Sana Jabeen SJ122 (LAH35077).

Molecular phylogenetic characterization (Figures 89 & 90)

Sequencing of the PCR products of ITS region of Verpa asiatica yielded 870–889 base pairs by using ITS1F and ITS4 primers. A consensus sequence of 889 base pairs was BLAST searched at NCBI. It showed 99% identity to Verpa sp. from India (GQ281276, GQ281277 & JQ723135) and 90% to V. bohemica (Krombh.) J. Schröt. from Canada (GQ304945), Germany (AJ648479), Poland (AM269562) and Ukrine (JQ723135) 81– 92% query cover 0.0 E value.

Sequencing of the PCR products of LSU yielded 1154–1243 base pairs by using LR0R and LR5 primers. Consensus sequence of 1154 base pairs was BLAST searched at NCBI. The subject sequence showed 99% identity to V. bohemica from USA (KC751530

168

& FJ176853) and 97% identity to V. conica (O.F. Müll.) Sw. from Germany (AJ698470) and USA (KC751531 & AY544666) with 73–74% query cover and 0.0 E value.

To reconstruct phylogeny, closely related ITS and LSU sequences were retrieved from the GenBank. Taxa from genus Morchella Dill. ex Pers were chosen as out groups. The sequences generated during this study diverged from the other identified taxa with strong boot strap value in both ITS and LSU based sequence analysis.

Comments

Verpa asiatica is characterized by its free, brown hymenium bearing vertical ridges, off white to yellowish hollow hymenophore, asci with a long narrow base consisting of two oblong smooth spores, and frequently septate paraphyses. Morphologically the species is closely related to V. bohemica, but anatomically it can be distinguished from V. bohemica on the basis of spore morphology. The spores of V. bohemica are slightly curved or boat shaped with somewhat narrower endings (Healy et al., 2008), while the spores of V. asiatica have smooth elliptical spores with parallel side walls. Molecular data analysis based on ITS and LSU sequences also proved it a distinct taxon.

169

Figure. 87. Morphology of Verpa asiatica. Ascomata LAH35077 (holotype). Bar = 2 cm.

170

A

B C

Figure 88. Anatomy of Verpa asiatica. A–C. LAH35077. A. Ascospores; B. Paraphyses; C. Asci. Bars: A = 0.8 µm; B = 90 µm; C = 40 µm.

171

75 JF908754 Verpa conica

88 JQ723141 Verpa sp JQ723142 Verpa cf conica 100 JN043311 Verpa conica 73 JQ723143 Verpa sp JQ723145 Verpa sp 54 99 JQ723144 Verpa sp 93 AJ544205 Verpa conica Clade I JQ723140 Verpa sp 97 100 AF008230 Verpa conica AJ544206 Verpa conica 89 80 AJ544204 Verpa conica GQ223458 Uncultured Verpa JQ723147 Verpa sp JQ723146 Verpa sp 100 JQ723148 Verpa sp Verpa asiatica SJ120 50 JQ723135 Verpa sp

100 Verpa asiatica SJ122 GQ281276 Verpa sp GQ281277 Verpa sp 65 AM269502 100 AJ698479 Verpa bohemica Clade II JQ723138 Verpa sp 96 100 GQ304945 Verpa bohemica JQ723139 Verpa sp JQ723137 Verpa bohemica 23 JQ691487 Verpa bohemica JQ723136 Verpa bohemica GQ325237 Out group

0.05 Figure 89. Molecular phylogenetic analysis of Verpa asiatica based on ITS sequences. The evolutionary history was inferred by the Maximum Likelihood method based on Jukes-Cantor model. The analysis involved 30 nucleotide sequences. There were a total of 939 positions in the final dataset. Sequences generated during this study are marked with .

172

KC751531 Verpa conica

94 AY544666 Verpa conica

66 AJ698470 Verpa conica

Verpa asiatica SJ120

100 KC751530 Verpa bohemica 52

97 FJ176853 Verpa bohemica

AJ698472 venosa

88 AY544667 56 92 KC751497 Disciotis venosa

GQ119355 carthusianum 84 GQ119357 Leucangium carthusianum

100 GQ119359 Leucangium sp.

77 GQ119358 Leucangium sp

EU327203 gigantea

100 EU327201 Imaia gigantea

96 EU327202 Imaia gigantea

AF279398 Morchella esculenta Out group

0.01 Figure 90. Molecular phylogenetic analysis of Verpa asiatica based on LSU sequences. The evolutionary history was inferred by the Maximum Likelihood method based on Jukes-Cantor model. The analysis involved 17 nucleotide sequences. There were a total of 711 positions in the final dataset. Sequence generated from LAH35076 are marked with .

173

Genus Peziza

Peziza khanspurensis nom. prov. (Figures 91 & 92)

Etymology: The specific epithet refers to Khanspur, the site of collection.

Ascomata upto 1.3 cm high, up to 4 cm wide; cup shaped to irregular, brittle, margins undulating, hymenium up to 0.25 cm thick, surface granulose to verrucose; sessile; light brown (2.5Y7/6) to grayish brown (10YR3/4) or dark brown (10YR2/4) at maturity.

Ascospores [30/3/2] (16.1) 17.4–19.9 (20.2) × (9.7) 9.8–10.9 (12.1) µm, Q = (1.5) 1.6–1.9 (2), avQ = 1.81; smooth, subglobose to ellipsoid, apiculus invisible. Asci (266.4) 267–275.4 (283.2) × (12.5) 12.8–13.7 (14.2) µm, cylindrical becoming narrower towards the base, base 6.8–10.8 µm, 8 spored, hyaline in 5% KOH. Paraphyses long up to 167 µm; wide up to 6.9 µm, filamentous with slightly clavate apices; hyaline in 5% KOH.

Material examined: Pakistan, Khyber Pakhtunkhwa province, Abbottabad district, Khanspur, 2575 m asl, associated with Cedrus deodara (Roxb. ex D. Don) G. Don), 19 May 2014 Lubna Choudry and Abdul Nasir Khalid SJ97; ANP1(LAH35091); 20 May 2014 Sana Jabeen and Abdul Nasir Khalid SJ123 (LAH35096); SJ124 (LAH35097).

Molecular phylogenetic characterization (Figure 93)

Sequencing of the PCR products of ITS region of Peziza khanspurensis yielded 684–694 base pairs by using ITS1F and ITS4 primers. A consensus sequence of 600 base pairs was BLAST searched at NCBI. It showed 95% identity to P. varia (Hedw.) Alb. & Schwein. (AF491545– AF491549 & AY789392) from USA with 95% query cover and 0.0 E value.

To reconstruct phylogeny, closely related ITS sequences were retrieved from the GenBank. Geopora cercocarpi D. Southw. & J. L. Frank (NR_121491) was chosen as out group. The sequences from Pakistan clustered in clade 1 with P. varia with 78% bootstrap support. These sequences got separated from P. varia forming a sister lineage of its own with 97% boot strap support.

174

Comments

Peziza khanspurensis is characterized by its medium sized grayish brown to dark brown cup shaped to irregular, brittle acomata with undulating margins and granulose to verrucose surface, smooth, subglobose to ellipsoid ascospores, septate paraphyses with slightly septate apices. It differs from P. varia because of its darker granular inner surface which is smooth and brown, the outer surface is pale, often almost white (Fries, 1822; Hansen et al., 2002). Peziza varia collections as studied by Hansen et al. (2002) on the basis of its smooth spores. Two lineages identified by them indicating two distinct species on the basis of spore ornamentation. Peziza khanspurensis also differs from these taxa because of its smooth spores. The species formed its own lineage by the separation from P. varia representatives with low warts.

175

Figure. 91. Morphology of Peziza khanspurensis. A & B. Ascomata A. LAH35091 (holotype); B. LAH35096. Bar = 0.5 cm.

176

A

B C

Figure 92. Anatomy of Peziza khanspurensis. A–C. LAH35091. A. Ascospores; B. Paraphyses; C. Asci. Bars: A = 13 µm , B = 0.2 µm, C = 32 µm.

177

97 Peziza khanspurensis SJ97 Peziza khanspurensis SJ124 97 123 AF491548_Peziza_varia_ AF491549_Peziza_varia AF491547_Peziza_varia 58 99 AF491546_Peziza_varia AF491545_Peziza_varia AF491551_Peziza_varia AF491550_Peziza_varia_ 82 AF491552_Peziza_varia 95 AF491554_Peziza_varia 60 AF491553_Peziza_varia AF491560_Peziza_varia Clade I KU061019_Peziza_varia_ KC832904_Peziza_cf_domiciliana 78 97 JF908557_Peziza_varia AF491570_Peziza_varia 82 AF491565_Peziza_varia AF491568_Peziza_varia_ 95 FJ235144_Peziza_cf_echinispora FJ235142_Peziza_cf_echinispora_ 99 AF491572_Peziza_sp_ 58 AF491571_Peziza_sp JF908533_Peziza_echinospora 34 AF491575_Peziza_echinispora_ 99 AF491574_Peziza_echinispora 90 AF491573_Peziza_echinispora AF491593_Peziza_fimeti 90 AF491597_Peziza_fimeti 22 AF491595_Peziza_fimeti_ JQ654493_Peziza_fimeti 86 AF491599_Peziza_fimeti 52 JQ654488_Peziza_fimeti AF491598_Peziza_fimeti 99 JF908549_Peziza_udicola 86 AF491609_Peziza_sp_ AF491616_Peziza_sp 99 AF491615_Peziza_sp 99 AF491587_Peziza_sp_ 64 AF491586_Peziza_sp_ Clade II 76 KC832902_Aleuria_amplissima_ 54 KC832901_Peziza_pseudovesiculosa 58 64 KC832903_Aleuria_amplissima AF491579_Peziza_arvernensis 92 AF491580_Peziza_arvernensis_ 80 KC832896_Peziza_arvernensis KC832897_Peziza_arvernensis_ JF908569_Peziza_arvernensis 93 KP125489_Peziza_arvernensis AF491583_Peziza_arvernensis 78 AF491581_Peziza_arvernensis AF491585_Peziza_arvernensis NR_121491_Geopora_cercocarpi Out group

0.05 Figure 93. Molecular phylogenetic analysis of Peziza khanspurensis based on ITS sequences. The evolutionary history was inferred by the Maximum Likelihood method using Jukes-Cantor model. The analysis involved 54 nucleotide sequences. There were a total of 755 positions in the final dataset. Sequences generated from Pakistani collections are marked with .

178

Peziza succosella (Le Gal & Romagn.) M.M. Moser ex Aviz.-Hersh. & Nemlich, Israel J. Bot. 23: 156 (1974)

Jabeen et al., 2016 (Annexture)

179

Genus Geopora

Geopora pinyonensis L. Flores-Rentería & C.A. Gehring, Mycologia 106: 556 (2014)

(Figures 94 & 95)

Ascomata up to 0.8 cm high, upto 1.3 cm wide, deeply cup shaped, margins slightly outwards, undulating and rimose, up tp 0.12 cm wide, submerged in the soil, sessile. Surface granulose, whitish (2.5GY9/2) to light brown (5Y8/4).

Ascospores [30/3/2] (24) 25.5–27.4 (28.9) × (12.8) 13.6–14.4 (15.3) µm, Q = (1.6) 1.7–2 (2.1), avQ = 1.9; smooth, subglobose to ellipsoid, apiculus in visible. Asci (228.6) 235.9–254 (256.7) × (15.5) 16.4–20.8 (21.7) µm, cylindrical having a narrow base , base up to 9.5 µm, 8 spored, biseriate spores also observed inside the asci,; hyaline in 5% KOH. Paraphyses 4.8–6.1 µm wide, up to 182.3 µm long, septate, filamentous, apices slightly clavate becoming narrow towards the base; hyaline in 5% KOH.

Material examined: Pakistan, Khyber Pakhtunkhwa province, Hazara division, Mansehra district, 2500 m asl, associated with Cedrus deodara (Roxb. ex D. Don) G. Don), 3 Aug 2014, Sana Jabeen and Abdul Nasir Khalid SJ139 (LAH35104); (LAH35105).

Molecular phylogenetic characterization (Figures 96)

Sequencing of the PCR products of ITS region of Geopora pinyonensis yielded 633–637 base pairs by using ITS1F and ITS4 primers. A consensus sequence of 637 base pairs was BLAST searched at NCBI. It showed 67–99% identity to G. arenicola (FM206449 & FM206453) from Estonia, G. cf. cervina (DQ200831) from Norway, G. pinyonensis (KF768652) and sp. (AF266709) from USA and Pyronemataceae sp. (GQ281481) from Germany.

To reconstruct phylogeny, closely related ITS sequences were retrieved from the GenBank. Tarzetta catinus (Holmsk.) Korf & J. K. Rogers (FM206478) was chosen as out group. The sequences from Pakistan clustered in a same clade with G. pinyonensis with strong boot strap support.

180

Comments

Geopora are important ectomycorrhizal associates that dominate the plant communities. Several members of the genus are known only from ectomycorrhizal tissue because of small and fairly hypogeous sporocarps. Geopora pinyonensis is a newly described taxon which was collected from northern Arizona, USA from under the pinyon (Pinus edulis Engelm.) (Flores-Rentería et al., 2014). In this study, the presence of G. pinyonensis extends its distribution to the Asian continent. It is the first report of this taxon from Pakistan.

181

Figure. 94. Morphology of Geopora pinyonensis. A & B. Ascomata A. LAH35104; B. LAH35105. Bars: A & B = 0.4 cm.

182

A

B C

Figure 95. Anatomy of Geopora pinyonensis. A–C. LAH35104. A. Ascospores; B. Paraphyses; C. Asci. Bars: A = 15 µm , B = 10 µm, C = 12.5 µm.

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86 JF908022_Geopora_nicaeensis 100 KT275618_Geopora_sp 100 KT275630_Geopora_sp FM206429_Geopora_sp 90 100 FM206430_Geopora_sp DQ200831_Geopora_cf_cervina 96 FM206427_Geopora_sp 100 FM206421_Geopora_sp FM206400_Geopora_tenuis 46 JQ724065_Geopora_cervina 100 94 FM206410_Geopora_cervina 58 100 KC840612_Cadophora_finlandica FM206432_Geopora_sepulta FM206431_Geopora_sepulta 50 100 KP745606_Geopora_sp AF266709_Pezizales_sp GQ281481_Pyronemataceae_sp 50 KF768652_Geopora_pinyonensis 100 Geopora pinyonensis SA139b 56 98 78 Geopora pinyonensis SA139a FM206458_Geopora_arenicola 84 FM206441_Geopora_arenicola FM206451_Geopora_arenicola FM206449_Geopora_arenicola FM206448_Geopora_arenicola 84 96 FM206446_Geopora_arenicola FM206445_Geopora_arenicola

32 FM206460_Geopora_arenicola FM206437_Geopora_arenicola 76 FM206452_Geopora_arenicola FM206439_Geopora_arenicola 94 FM206453_Geopora_arenicola FM206433_Geopora_arenicola HQ283092_Geopora_cercocarpi HQ283094_Geopora_cercocarpi NR_121491_Geopora_cercocarpi 100 HQ283093_Geopora_cercocarpi HQ283090_Geopora_cercocarpi FM206478_Tarzetta Out group

0.05 Figure 96. Molecular phylogenetic analysis of Geopora pinyonensis based on ITS sequences. The evolutionary history was inferred by the Maximum Likelihood method based on Jukes-Cantor model. The analysis involved 39 nucleotide sequences. There were a total of 646 positions in the final dataset. Sequences generated during this study are marked with .

184

4.2. IDENTIFICATION OF ECTOMYCORRHIZAL HOST

Cedrus deodara (Roxb. ex D. Don) G. Don., Hort. Brit. 1: 388 ( 1830)

Molecular phylogenetic characterization (Figure 97)

Sequencing of the PCR products of ITS region of 28S rRNA from the ectomycorrhizal tissue using 28KJ and 28C as forward and reverse primers respectively, yielded up to 1396 base pairs of nucleotides. Consensus sequences were obtained by the sequences from both the primers of individual sample were BLAST searched at NCBI. These sequences showed 96–99% identity to Abies fabri (Mast.) Craib (EU161347) and Cedrus atlantica (Endl.) Manetti ex Carrière (EU161348) from China, C. libani A. Rich. (AY056507) from Sweden, Pinus embra L. (U90681) and P. strobus L. from France and Sweden respectively with 46–96% query cover and 0.0 E value.

To reconstruct phylogeny, closely related sequences were retrieved from the GenBank. Ginko biloba L. (AY095475) and Pseudotaxus chienii (W. C. Cheng) W. C. Cheng (EU161465) were chosen as out groups. The sequences generated during this study clustered with the similar taxa. Sequences of C. deodara from Pakistan clustered with C. atlantica and C. libani with 100% bootstrap value. Abies sp. and Pinus spp. from Pakistan clustered in their respective clades with strong bootstrap value.

Comments

Cedrus deodara is an accepted species native to Himalayan region. It is a high mountain tree occurring in a wide range of habitats in the Himalaya. Several synonyms has been used for the species. It has been known as Abies deodara (Roxb. ex D.Don) Lindl., C. indica Chambray, Larix deodara (Roxb. ex D. Don) K. Koch and Pinus deodara Roxb. ex D. Don. (http://www.theplantlist.org/).

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AY056499 Pinus peuce AY056501 Pinus strobus 96 Pinus sp SA109

100 Pinus sp SA173 64 Pinus sp SA335 AY056500 Pinus mugo 88 72 EU161304 Pinus massoniana AY056509 Picea asperata AJ271027 Sequoiadendron giganteum 100 EU161351 Picea smithiana 50 AY056510 Picea breweriana 98 AY056511 Tsuga canadensis 94 EU161350 Tsuga dumosa EU161313 Nothotsuga longib

68 EU161305 Pseudolarix amabi EU161359 19661 Keteleeria davidi 32 EU161347 Abies fabri 46 Abies sp SA375 98 AY056508 Abies grandis AY056507 Cedrus libani Cedrus deodara SA139 62 Cedrus deodara SA201 Cedrus deodara SA208 100 EU161348 Cedrus atlantica Cedrus deodara SA27

52 Cedrus deodara SA25 Cedrus deodara SA207 Cedrus deodara SA333 Cedrus deodara SA332 EU161465 Pseudotaxus chienii Out group 100 AY095475 Ginkgo biloba

0.01 Figure 97. Molecular phylogenetic analysis of ectomycorrhizal hosts based on ITS sequences. The evolutionary history was inferred by the Maximum Likelihood method based on Tamura-Nei model. The analysis involved 31 nucleotide sequences. There were a total of 652 positions in the final dataset. Sequences generated during this study are marked with .

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4.3. CHARACTERIZATION OF NON ECTOMYCORRHIZAL FRUIT BODIES Table 2. Non ectomycorrhizal fruit bodies collected from different stands

Collection GenBank BLAST Query Level of Taxon Voucher Region/Gene Identity Number Accessions Match Cover Identification Agaricus augustus Fr. SJ107; MTD LAH35093 ITS - JF797193 100% 99% Species SJ60; K4-14 LAH35085 ITS - AY899266 98% 98% Agaricus sp. 1 ITS - AY899266 98% 98% Genus SJ53; Mt2-1 LAH35084 LSU - KP331529 74% 99% Armillaria sp. 1 SJ83 K4 LAH35088 ITS - KF114476 100% 96% Genus Calocera sp.1 SJ127; MP LAH35098 ITS - KC581302 46% 94% Genus SJ93; MP8 LAH35089 ITS - JX968260 99% 89% Conocybe sp. 1 Genus SJ96; MP4 LAH35090 ITS - JX968360 99% 94% asperum SJ46; K1 LAH35082 ITS - KC581336 100% 99% Species (Pers.) Bon Gerhardtia borealis (Fr.) Contu & A. SJ28; K4b LAH35080 ITS - KR673544 97% 99% Species Ortega Gymnopilus sp. 1 SJ108; 19M LAH35094 ITS - KR072502 74% 97% Genus Gymnopus sp. 1 SJ81; K4 LAH35087 ITS - AM505783 95% 90% Genus cristata SJ104; Kh-1 LAH35092 ITS - EU081960 100% 99% Species (Bolton) P. Kumm.

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Lepiota sp. 1 SJ71; K19 LAH35086 ITS - KC556779 100% 98% Genus KU647726, ITS KU647727, JX896447 100% 93% Leucoagaricus KU647728 pakistaniensis nom. SJ13; K4-A LAH35013 LSU - KF571951 99% 98% Species prov. KU900514, rpb2 HM488857 83% 95% KU900515 tef-1α KU900512 HM488913 90% 91% Macrolepiota LAH-SJ1- excoriata (Schaeff.) SJ2; K3-1 ITS KJ013326 JQ683100 100% 99% Species 2013 M.M. Moser Melanoluca sp. 1 SJ52; K4-31 LAH35083 ITS - JF908357 90% 98% Genus rosella (Fr.) SJ15; KU-5 LAH35079 ITS - JF908473 100% 99% Species P. Kumm. Pluteus pouzarianus SJ12; KLM LAH35078 ITS - HM562096 99% 99% Species Singer Psathyrella candolleana (Fr.) SJ114; D LAH35095 ITS - EU520251 98% 99% Species Maire Scutellinia crinita SJ133; HP LAH35100 ITS - AY220799 90% 97% Species (Bull.) Lambotte SJ137; HP2 LAH35102 ITS - DQ491492 52% 89% Scutellinia sp. 1 Genus SJ138; HP LAH35103 ITS - DQ491492 52% 89% Xeromphalina sp. 1 SJ37; K2/1 LAH35081 ITS - GU320004 64% 99% Genus Xeromphalina sp. 2 SJ129 LAH35099 ITS - KM024581 81% 90% Genus Xylaria psidii J. D. SJ135; K4 LAH35101 ITS - KJ767111 96% 99% Species Rogers & Hemmes

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4.4. CHARACTERIZATION OF SAPROBIC FUNGI FOUND FROM FRUIT BODIES

Table 3. Saprobic fungi found from fruit bodies

Collection GenBank BLAST Query Level of Taxon Region/Gene Identity Number Accessions Match Cover Identification penicillioides F134 ITS - KP131612 76% 86% Species Speg. Epicoccum nigrum Link F95 ITS - KT898754 78% 76% Species Eurotium sp. F128 ITS - KF639819 87% 84% Genus

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4.5. CHARACTERIZATION OF BELOW GROUND ECTOMYCORRHIZAL FUNGI

Table 4. List of below ground ectomycorrhizal fungi/OTUs

GenBank BLAST Query Level of Taxon Collection Number Voucher Identity Accessions Match Cover Identification SA205; KH-3 LAH-EM8-2013 - HG796868 77% 99% Amphinema sp. 1 Genus SA322; MK3c LAH-EM9-2013 - HG796872 66% 93% Clavulicium delectabile (H.S. SA65; K4-2MT LAH-EM10-2013 - KP783435 90% 97% Species Jacks.) Hjortstam cf. cinerea SA224; KF6a LAH-EM11-2013 - EU862214 93% 98% Species SA11; K4-19 LAH-EM12-2013 - JF273520 60% 87% Clavulina sp. 1 SA331; KF3 LAH-EM13-2013 - KC684922 85% 98% Genus SA333b; KF2b LAH-EM14-2013 - KC684922 76% 98% Clavulina sp. 2 SA28; MT LAH-EM15-2013 - KT447177 95% 99% Genus Cortinarius leucopus SA206a; Hp-14-4a LAH-EM16-2013 - JN133921 94% 98% Species (Bull.) Fr. Cortinarius oulankaensis Kytöv., SA70; K4-12b LAH-EM17-2013 - FJ039672 86% 98% Species Niskanen, Liimat. & H. Lindstr. Inocybe oblectabilis SA342; KF11 LAH-EM18-2013 - JF927857 85% 98% Species (Britzelm.) Sacc. SA214; KF9 LAH-EM19-2013 - JF927857 86% 99%

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SA2; K4-18 LAH-EM20-2013 - KM576451 84% 93% Inocybe sp. 1 Genus SA36; K LAH-EM21-2013 - KM576451 88% 93% SA112;HP-7b LAH-EM22-2013 - KM576451 62% 93%

SA336a; MK2 LAH-EM23-2013 - KM576451 83% 92% Inocybe sp. 2 SA23; K2-6 LAH-EM24-2013 - HG796986 93% 99% Genus Inocybe sp. 3 SA43; MT2-4 LAH-EM25-2013 - HQ604085 53% 95% Genus Inocybe sp. 4 SA69; K4-12a LAH-EM26-2013 - HG796980 91% 99% Genus SA69Z; K4-12c LAH-EM27-2013 - HG796986 89% 99% SA71; K4-12d LAH-EM28-2013 - HG796986 93% 99% Genus Inocybe sp. 5 SA86; K4-8 MT LAH-EM29-2013 - HG796986 92% 99%

SA98; K4-1 MTb LAH-EM30-2013 - HG796986 89% 99% SA111; HP-7 LAH-EM31-2013 - HG796986 88% 99% SA76; K4-18 LAH-EM31-2013 - HG796992 88% 93% Inocybe sp. 6 SA148; KG-9 LAH-EM32-2013 - HG796992 87% 99% Genus SA369; SK6b1 LAH-EM33-2013 - HG796992 81% 96% SA120; HP-52 a LAH-EM34-2013 - KF679813 90% 99% Inocybe sp. 7 Genus SA123; HP52-c LAH-EM35-2013 - KF679813 97% 98% Inocybe sp. 8 SA165; Pt-7 LAH-EM36-2013 - HG796989 85% 96% Genus Inocybe sp. 9 SA201; Kh-4 LAH-EM37-2013 - JF908235 73% 97% Genus

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Inocybe sp. 10 SA203; Kh2-a LAH-EM38-2013 - AM882932 91% 94% Genus Inocybe sp. 11 SA208; Kh-15a LAH-EM39-2013 - HQ604207 86% 85% Genus Inocybe sp. 12 SA216; KF20a LAH-EM40-2013 - HG796964 85% 95% Genus SA315; MK1a LAH-EM41-2013 - HE862959 92% 99% Genus SA373; MJ3 LAH-EM42-2013 - HG796969 60% 96% Genus Inocybe sp. 13 SA325a; KF12 LAH-EM43-2013 - JF908187 78% 89% Genus SA315; MK1b LAH-EM44-2013 - HE862959 78% 99% Genus Inocybe sp. 14. SA375; T62 LAH-EM45-2013 - AM882875 55% 81% Genus Peziza succosella (Le Gal & Romagn.) M.M. SA116; HP-5 LAH-EM1-2013 KM199728 KP311465 88% 99% Species Moser ex Aviz.-Hersh. & Nemlich Piloderma sp. 1 SA324a; KF-16 LAH-EM46-2013 - KP814544 98% 98% Genus Russula amethystina SA217; KF6b2 LAH-EM47-2013 - KF679818 87% 99% Species Quél. Russula anthracina SA99; K4-5 LAH-EM2-2013 KR011881 KF679819 85% 99% Species Romagn.

SA317; HP-14-3 LAH-EM7-2014 KT944027 95% 99% Species JN129398 Russula livescens (Batsch) Bataille SA206b; HP-14-4b LAH-EM6-2014 KT944026 JN129398 94% 99% Species SA107; HP-6 LAH-EM4-2013 KR020028 JN129398 93% 99% Genus

Russula pakistanica SA48; K4-100a LAH-EM48-2013 - HG796943 95% 86% Species nom. prov. SA110; Pt LAH-EM49-2013 - HG796943 94% 98%

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SA158; Pt-4 LAH-EM50-2013 - HG796943 91% 96% SA166; Pt-7b LAH-EM51-2013 - HG796943 92% 96% SA171; Pt-3c LAH-EM3-2013 KT834642 HG796943 88% 99% SA178; HP-14-3c LAH-EM5-2013 KT944025 JN129398 96% 99% Russula sp. 1 SA55a; K2-20 LAH-EM52-2013 - KR673656 80% 81% Genus sp. 1 SA329b; MK3b LAH-EM53-2013 - HQ215831 88% 99% Genus Tomentella sp. 1 SA119; Hp-5d LAH-EM54-2013 - AY635176 93% 99% Genus Tomentella sp. 2 SA348; KF1 LAH-EM55-2013 - HQ215824 87% 96% Genus Tomentella sp. 3 SA374; C1K LAH-EM56-2013 - JQ711817 82% 96% Genus Tricholoma aurantium SA213; KF8 LAH-EM57-2013 - DQ367919 98% 99% Species (Schaeff.) Ricken Trichophaea sp. 1 SA208b; KH-15a LAH-EM58-2013 - JN129415 92% 99% Genus SA3; K4-18b LAH-EM59-2013 - KC517481 93% 99% Genus Tuber sp. 1 SA106; Pt-19 LAH-EM60-2013 - KC517481 93% 98% Genus Tuber sp. 2 SA206a; Hp-14-4d LAH-EM61-2013 - GU979046 95% 80% Genus SA37; MT-12 LAH-EM62-2013 - JX129137 73% 89% Genus Wilcoxina sp. 1 SA103; MT-20 LAH-EM63-2013 - JX129137 45% 75% Genus

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4.6. CHARACTERIZATION OF BELOW GROUND NON ECTOMYCORRHIZAL FUNGI Table 5. List of below ground non ectomycorrhizal fungi

Sample BLAST Query Level of Taxon Identity Number Match Cover Identification Aspergillus protuberus Munt.-Cvetk. SA218 NR_135353 83% 100% Species Byssochlamys verrucosa Samson & Tansey SA349 KM519668 88% 99% Species Coniochaeta mutabilis (J.F.H. Beyma) Z.U. Khan, SA223 KM056320 85% 94% Species Gené & Guarro Galactomyces geotrichum (E.E. Butler & L.J. Petersen) SA226b JN903644 80% 99% Species Redhead & Malloch Geomyces sp. SA203 KP714581 75% 99% Species Gymnostellatospora alpina (E. Müll. & Arx) Udagawa SA333c FJ590609 73% 95% Species Gymnostellatospora alpina SA331b FJ590609 74% 99% Species Lecythophora sp. SA329 KJ957772 93% 98% Species Lecythophora sp. SA212 KJ957772 70% 97% Species Lecythophora sp. SA318 KC007198 93% 100% Species Lecythophora sp. SA325 KJ957772.1 85% 98% Species Microdochium sp. 1 SA326b AM502266 81% 92% Genus

Mortierella sp. 1 SA130 KP311420 64% 99% Genus Penicillium sp. 1 SA130 HQ850339 82% 98% Genus Phialocephala fortinii C.J.K. Wang & H.E. Wilcox SA56 JQ711965 82% 98% Species

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Phialocephala fortinii SA213 JQ711965 87% 99% Species Phialocephala fortinii 216b JQ711965 81% 97% Species Phialocephala fortinii SA224b JQ711965 90% 99% Species Phialocephala fortinii SA332 JQ711965 93% 99% Species Phialocephala fortinii SA333a AY524846 35% 97% Species Phialocephala fortinii SA343 JQ711965 94% 99% Species Phialocephala fortinii SA345 AY394921 96% 99% Species Phialocephala helvetica Grünig & T.N. Sieber SA211 EU103612 89% 97% Species Phialocephala helvetica SA335 EU103612 94% 94% Species Phialocephala helvetica SA336b EU103612 86% 83% Species Phialocephala helvetica SA340 EU103612 83% 100% Species Phialocephala helvetica SA339 EU103612 83% 99% Species Phialocephala sp. 1 SA326a AB671499.2 83% 82% Genus sp. 1 SA300 KM576578 76% 98% Species Trichocladium sp. 1 SA207 AM292049 94% 95% Genus Trichocladium sp.1 SA324b AM292049 94% 95% Genus Trichoderma crassum Bissett SA26 NR_134370 90% 100% Species Trichoderma sp. 1 SA55b; K2-20 NR_134362 36% 78% Genus Xylogone sphaerospora Arx & T. Nilsson SA27 JN198528 82% 99% Species

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4.7. IDENTIFICATION OF HOST TAXA FROM ECTOMYCORRHIZAL MORPHOTYPES

Table 6. Host tree species identified from ectomycorrhizal morphotypes

BLAST Query Level of Taxon Collection Number Identity Match Cover Identification EU161347 Abies sp. T62; SA375 95% 99% Genus

KG-9; SA148 EU161348 95% 99% Species EU161348 KH-4; SA201 56% 99% Species

SA204 EU161348 95% 99% Species SA207 EU161348 96% 99% Species KH-15a; SA208 EU161348 68% 99% Species KF8; SA213 EU161348 46% 99% Species Cedrus deodara SA224 EU161348 47% 99% Species SA229 EU161348 60% 99% Species SA331 EU161348 65% 99% Species SA332 EU161348 65% 99% Species SA333 EU161348 96% 99% Species SA334 EU161348 78% 97% Species

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SA139 EU161348 91% 99% Species SA152 EU161348 96% 99% Species SA226 EU161348 69% 99% Species KF6b EU161348 96% 99% Species 17 EU161348 96% 99% Species 25 EU161348 95% 99% Species 26 EU161348 85% 98% Species Cedrus deodara 43 EU161348 95% 99% Species

SA27 EU161348 88% 99% Species SA29 AY056507 96% 96% Species KF20a; SA216 AY056507 73% 86% Species SA330 AY056507 80% 99% Species MJ3; SA373 U90681 81% 99% Genus HP-3; SA103 AY056501 87% 99% Genus Pt-18; SA109 AY056501 89% 99% Genus Pinus spp. Pt-29; SA174 AY056501 72% 99% Genus KH-9; SA197 AY056501 87% 99% Genus HP-13a; SA198 AY056501 71% 99% Genus MT2; SA314 AY056501 61% 99% Genus

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MK4; SA335 AY056501 90% 99% Genus MJ4; SA368 AY056501 96% 99% Genus HP-9; SA173 AY056501 94% 99% Genus MK7; SA304 AY056501 96% 99% Genus Pt-17; SA134 AY056501 94% 99% Genus Pinus spp. Pt-21; SA142 AY056501 70% 97% Genus HP-52a; SA122 AY056501 94% 99% Genus Pt-13; SA129 AY056501 96% 99% Genus

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4.8. SOIL ANALYSIS

Table 7. Stand wise results of different soil analysis parameters

Hazara division Malakand division Rawalpindi division Parameters Stand 1 Stand 2 Stand 3 Stand 4 Stand 5 Stand 6 EC (mScm-1) 3.6 2.8 2 2 2.8 4.7 pH 6.6 6.8 6.1 6.45 6.3 5.8 Organic Matter (%) 0.91 0.87 0.85 0.83 0.81 0.89 Available Phosphorous (mg kg-1) 4.5 4 6.7 5.6 5.4 4.8 Available Potassium (mg kg-1) 118 110 130 114 102 118 Saturation (%) 42 38 38 40 40 42 Texture Loam Loam Loam Loam Loam Loam Boron (ppm) 1.45 1.45 1.13 1.13 1.08 1.08 Sulphate (meqL-1) 1.82 1.82 1.98 1.98 1.55 1.55 Zn (ppm) 2.5 2.5 2.14 2.14 1.89 1.89 Fe (ppm) 6.2 6.2 6.4 6.4 7.8 7.8 Cu (ppm) 1.27 1.27 0.47 0.47 0.44 0.44 Mn (ppm) 1.52 1.52 1.67 1.67 2.57 2.57 Ca + Mg (ppm) 2.8 1.8 1.4 1.5 2 4 Na (ppm) 8 10 6 5 8 7 SAR 2.1 3.3 2.2 1.8 2.5 1.5

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4.9. COMMUNITY ANALYSIS

A total of 107 fungal taxa were identified belonging to 67 genera and 50 families. Among these, 69 species belonging to 33 genera and 25 families were found to be ectomycorrhizal. From these 69 species, 33 species were identified as ectomycorrhizal fungal fruit bodies belonging to 19 genera and 14 families and 36 species belonging to 14 genera and 11 families were identified from ectomycorrhizal morphotypes. From the above and below ground data, four species were found as fruit bodies as well as in the form of ectomycorrhizae. So, 65 distinct ectomycorrhizal species were identified and remaining taxa belong to non ectomycorrhizal fungi and excluded in the final dataset for community analysis.

4.9.1. Alpha diversity

A total of 31 ectomycorrhizal fruit bodies were collected from Khanian forest (stand 1) represented by four taxa. These taxa include Cortinarius longistipus, Geopora pinyonensis, Inocybe alba. and I. flavellorimosa. Geopora pinyonensis was found most abundant from this site covering 80.64% of the community. Only one representative of Cortinarius longistipus was found with a relative abundance of 13.22%. Inocybe represented by two species; Inocybe alba and I. flavellorimosa were found to be the diverse genus in the form of basidiomata. Three individuals of I. alba and two of I. flavellorimosa were collected from this area with a relative abundance of 9.67 and 6.45%, respectively (Figures 98 & 99). Inocybe spp. and Corinarius longistipus were found to be the rare species in rank abundance curve while Geopora was dominating taxon in the stand (Figure 100). The median range (second quartile) of the abundance was 3.5 with 1 and 11 whiskers at the lowest and highest value, the abundance of each taxon in first and third quartile was 2.25 and 8.5, respectively showing large variations in the population size of the taxa in stand 1(Figure 131). The diversity index of stand 1 was calculated as 0.6538 based on Simpson's diversity index. Dominance index, Margalef's richness index and equitability index were calculated as 0.3462, 0.8736 and 0.4956, respectively (Table 8).

A total of 54 ectomycorrhizal morphotypes were identified from stand 1 belonging to two genera and six species namely Inocybe sp. 9, I. sp. 10, I. sp. 11, I. sp. 13, I. sp. 14 and

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Trichophea sp. 1. Inocybe sp. 13 was found most abundant represented by 15 morphotypes covering 23.43% of the community. Inocybe sp. 10 was represented by 11 morphotypes with 17.10% relative abundance. Inocybe sp. 9 was represented by 8 individuals with a relative abundance of 12.50%. Inocybe sp. 11 and Trichophea sp. 1 were represented by seven individuals of each showing 10.93% relative abundance. Inocybe sp. 14 was represented by six individuals with a relative abundance of 9.3% (Figure 101 & 102). All the species were found from only one stand and ranked at 1 as singletons in rank abundance curves (Figure 103). The median of the abundance was 8 with 6 and 15 whiskers at the lowest and highest value. The abundance of taxa in first and third quartile was 7 and 10.5, respectively (Figure 132). Simpsons diversity index was calculated as 0.1712. Dominance, Margalef's richness and equitability indices were calculated as 0.8288, 1.253 and 0.9692, respectively (Table 8).

From Khanspur (stand 2), 10 ectomycorrhizal ascomata were collected from under cedar belonging to four taxa viz; Gyromitra khanspurensis, Morchella pakistanica, Peziza khanspurensis and Verpa asiatica. Gyromitra khanspurensis was represented by two individuals with 20% relative abundance. Verpa asiatica was found most abundant, with four individuals representing 40% relative abundance. Three representatives of P. khanspurensis were collected from stand 2 covering the 30% of the community while only one specimen of M. pakistanica was collected from this site with a relative abundance of 10% (Figures 104 & 105). Peziza khanspurensis and V. asiatica were doubletons while G. khanspurensis and M. pakistanica were the singletons and regarded as rare species (Figure 106). The median of the abundance was 1.5 with 1 and 3 whiskers at the lowest and highest value. The abundance of taxa in first and third quartile was 2 and 4, respectively (Figure 131). Simpsons diversity index of stand 2 was calculated as 0.2222. Dominance, Margalef's richness and equitability indices were calculated as 0.7778, 1.303 and 0.9232, respectively (Table 8).

A total of 205 ectomycorrhizal morphotypes were identified from stand 2 belonging to six genera and nine species. The species include; Cortinarius leucopus, Inocybe sp. 1, I. sp. 5, I. sp. 7, Peziza succosella, R. livescens, R. pakistanica, Tomentella sp. and Tuber sp. 2. Russula livesence was found most abundant represented by 46 morphotypes covering 22.43% of the community. Peziza succosella was represented by 44 morphotypes with 21.46% relative abundance. Inocybe sp. 7 was represented by 31 morphotypes with a relative

201 abundance of 15.12%. Inocybe sp. 1 was represented by 23 morphotypes covering 11.11% of the community while 21 morphotypes of C. leucopus were found showing 10.24% relative abundance and 17 morphotypes fall in Inocybe sp. 5 with 8.29% relative abundance. Russula pakistanica and Tuber sp. 2 were represented by 9 morphotypes each with 4.39% abundance. Tomentella sp. covered 2.43% of the community with 5 morphotypes (Figures 107 & 108). All the species were found only once and regarded as rare species accept R. livescens and I. sp. 7. Russula livesence was found dominant taxon collected from three samples while I. sp. 7 was found as doubleton (Figure 109). The median of the abundance was 12 with 3 and 44 whiskers at the lowest and highest value. The abundance of taxa in first and third quartile was 9 and 21.5, respectively (Figure 132). Simpsons diversity index was calculated as 0.1495. Dominance, Margalef's richness and equitability indices were calculated as 0.8505, 1.503 and 0.911, respectively (Table 8).

From Kalam (stand 3), 104 ectomycorrhizal basidiomata were collected from under the cedar belonging to 9 genera and 15 species. These species include; Amanita brunneopantherina, A. flavipes, Cortinarius corrosus, Gomphidius flavostipus, Hebeloma angustisporium, Inocybe alba, I. mimica, I. oblectabilis, Neoboletus luridiformis, Russula anthracina, R. delica, R. amethystina, R. pakistanica, Suillus convexatus and Xerocomellus rimosus. Russula anthracina was the leading dominant with 26 individuals showing 25% relative abundance. Russula delica also covered a big part of the community with 25 individuals representing 24.04% relative abundance. Hebeloma angustisporium and I. mimica were represented by 7 individuals with an equal relative abundance of 6.73%. Inocybe oblectabilis and R. amethystina were represented by six individuals. Each covering the 5.77% of the community. Amanita flavipes, C. corrosus and G. flavostipus were represented by four individuals of each with an equal relative abundance of 3.85%. Only two individuals of Russula pakistanica and one of I. alba were found from with a relative abundance of 1.92 and 0.96, respectively (Figures 110 & 111). Amanita brunneopantherina, C. corrosus, H. angustisporium, I. alba, , S. convexatus and X. rimosus were singletons and represented the rare species, while A. flavipes, G. flavostipus, N. luridiformis and R. pakistanica were found as doubletons. Russula anthracina stood dominant taxon found to occur at rank 6 in abundance (Figure 112). The median of the abundance was given as 3 with 1 and 7 whiskers at the lowest and highest value. The abundance of taxa in first and third

202 quartile was 2 and 4, respectively (Figure 131). Simpsons diversity index of stand 2 was calculated as 0.1359. Dominance, Margalef's richness and equitability indices were calculated as 0.8641, 3.0140 and 0.8447, respectively (Table 8).

From this stand, 192 ectomycorrhizal morphotypes were identified. These morphotypes belong to six genera and 12 species. The species include; Clavulina sp. 1, C. cf. cinerea, I. oblectabilis¸ I. sp. 1, I. sp. 2, I. sp. 6, Piloderma sp. 1, Russula amethystina, R. sp. 1, Tomentella sp. 2, T. sp. 3 and Tricholoma aurantium. Inocybe sp. 2 was found as most abundant taxon with 46 morphotypes covering 23.96% of the community followed by T. aurantium represented by 31 morphotypes showing 16.15% relative abundance. Russula sp. 1, R. amethystina, Clavulina sp. 1, I. sp. 1, C. cf. cinerea, I. sp. 6, Tomentella sp. 3, I. oblectabilis, Piloderma sp. 1 and Tomentella sp. 2 were represented by 27, 22, 15, 14, 12, 8, 7, 5, 3 and 2 morphotypes showing 14.06, 11.46, 7.81, 7.29, 6.25, 4.17, 6.64, 2.60, 1.56 and 1.04% relative abundance, respectively (Figure 113 & 114). All the species were found from only one sample and ranked at 1 as singletons in rank abundance curves accept C. sp. 1, which was found from two samples and ranked as doubleton (Figure 115). The median of the abundance of taxa was 10 with 2 and 46 whiskers at the lowest and highest value. The abundance of taxa in first and third quartile was 5 and 22, respectively (Figure 132). Simpsons diversity index was calculated as 0.1313. Dominance, Margalef's richness and equitability indices were calculated as 0.8687, 2.092 and 0.8793, respectively (Table 8).

A total of 110 ectomycorrhizal fruit bodies were collected from Mashkun (stand 4) represented by 10 genera and 23 species. These species include Amanita ahmadii, A. brunneopantherina, A. flavipes, A. glarea, A. swatica, Boletus himalayensis, Cortinarius corrosus, C. longistipus, Gomphidius flavostipus, Hortiboletus rubellus, Inocybe alba, I. flavellorimosa, I. kohistanensis, I. mimica, I. sp. 15, Neoboletus luridiformis, Rhozopogon flavus, Russula amethystina, R. anthracina, R. delica, R. pakistanica, R. rubecola and Xerocomellus rimosus. Neoboletus luridiformis was found as most abundant taxon in the community represented by 18 individuals with a relative abundance of 16.36%. Amanita flavipes was second most abundant taxon represented by 15 individuals with 13.64% relative abundance followed by R. amethystina with 11 individuals covering 10% of the community. Rhizopogon flavus and I. kohistanensis were represented by seven and six individuals with a

203 relative abundance of 6.36 and 5.45%, respectively. Amanita swatica, C. corrosus and Russula pakistanica were represented by 5 individuals each with relative abundance of 4.54%. Cortinarius longistipus, G. flavostipus, I. mimica, R. anthracina, R. delica showed 3.64% relative abundance with 4 individuals of each taxon. Amanita ahmadii and A. brunneopantherina were equally represented by 3 individuals with 2.72% relative abundance of each. Xerocomellus rimosus was represented by two individuals covering 1.81% of the community. Amanita glarea, H. rubellus, I. alba, I. flavellorimosa, I. sp. 15 and R. rubecola were represented by only one individual in the stand with 0.90% relative abundance (Figures 116 & 117). Amanita ahmadii, A. glarea, C. corrosus, C. longistipus, H. rubellus, I. alba, I. flavellorimosa, I. sp. 15, Rhizopogon flavus, Russula rubecola and Xerocomellus rimosus were found only once and ranked as singletons. Amanita brunneopantherina, B. himalayensis, Gomphidius flavostipus, I. mimica, R. anthracina, R. delica, R. pakistanica were ranked second and found as doubletons. Amanita flavipes, N. luridiformis and R. amethystina were found as dominating taxa collected from six sites (Figure 118). The median abundance was 2 with 1 and 7 whiskers at the lowest and highest value, the abundance of each taxon in first and third quartile was given as 1 and 3, respectively (Figure 131). The diversity of stand 1 was calculated as 0.0703 based on Simpson's diversity index. Dominance index, Margalef's richness index and equitability index were calculated as 0.9296, 4.68 and 0.8938, respectively (Table 8).

From stand 4, 342 ectomycorrhizal morphotypes were identified. These morphotypes belong to nine genera and 16 species. The species include; Amphinema sp. 1, Clavulicium delectabile, Clavulina sp. 1, C. sp. 2, Cortinarius oulankaensis, Inocybe sp. 1, I. sp. 13, I. sp. 3, I. sp. 4, I. sp. 5, I. sp. 6, Russula anthracina, R. pakistanica, Thelephora sp. 1, Tuber sp. 1and Wilcoxina sp. 1. Clavulicium delectabile was found as most abundant taxon with 68 morphotypes covering 19.88% of the community followed by I. sp. 5 represented by 46 morphotypes showing 13.45% relative abundance. Inocybe sp. 1, Cortinarius. oulankaensis, I. sp. 4, I. sp. 6, I. sp. 13, I. sp. 3, R. anthracina, Tuber. sp. 1, A. sp. 1, Thelephora sp. 1, W. sp. 1, R. pakistanica, Clavulina sp. 2 and C. sp. 1 were represented by 39, 30, 28, 20, 19, 19, 13, 11, 10, 10, 9, 8, 7 and 5 morphotypes showing 11.40, 8.77, 8.19, 5.85, 5.55, 5.55, 3.80, 3.21, 2.92, 2.92, 2.63, 2.33, 2.04 and 1.46% relative abundance, respectively (Figures 119 & 120). All the species were found from only one sample and ranked at 1 as singletons in rank

204 abundance curves accept I. sp. 1, I. sp. 13 and W. sp. 1, which were found from two samples and ranked as doubleton while I. sp. 5 was found from four samples and ranked 4 (Figure 121). The median of the abundance of taxa was 11 with 4 and 68 whiskers at the lowest and highest value. The abundance of taxa in first and third quartile was 8.5 and 17.75, respectively (Figure 132). Simpsons diversity index was calculated as 0.0980. Dominance, Margalef's richness and equitability indices were calculated as 0.902, 2.571 and 0.9055, respectively (Table 8). From Kuzah Gali (stand 5), 84 ectomycorrhizal morphotypes were identified. These morphotypes belong to Inocybe sp. 6 and covered the whole community and ranked 1 in species accumulation curve (Figures 125–127).

From Kuzah Gali (stand 5), seven basidiomata were collected representing four genera with one species of each. The species include; Geastrum galiyensis, Russula delica, R. pakistanica and Tricholoma terreum. Russula delica was found leading dominant represented by three individuals covering 42.86% of the community. Geastrum galiyensis was represented by two individuals with 28.57% relative abundance while one individual of R. pakistanica and T. terreum were found representing 14.28% relative abundance (Figures 122 & 123). Russula pakistanica and T. terreum were the rare species and found as singletons, G. galiyensis was found as doubleton and Russula delica was the most abundant and dominant species and ranked three in the rank abundance curve (Figure 124). In box plot, the median of the abundance of stand 5 was given as 1 with 1 and 2 whiskers at the lowest and highest value, the abundance of each taxon in first and third quartile was 1 and 2, respectively (Figure 131). The diversity of stand 5 was calculated as 0.1905 based on Simpson's diversity index. Dominance index, Margalef's richness index and equitability index were calculated as 0.8095, 1.542 and 0.9212, respectively. Additional parameters for alpha diversity analysis of the above ground taxa from all the stands are given in Table 8. . From stand 6 (Patriata), above ground representatives of the ectomycorrhizal fungi were not observed.

From Patriata stand 6, 42 ectomycorrhizal morphotypes were identified. These morphotypes belong to three genera represented by one species each. These species include; I. sp. 8, Russula pakistanica and Tuber sp. 1. Russula pakistanica was found most abundant taxon with 29 morphotypes covering 69.05% of the community followed by T. sp. 1 and I.

205 sp. 8 with nine and four morphotypes representing 21.42 and 9.52% of the community, respectively (Figure 128 & 129). Inocybe sp. 8 and T. sp. 1 were ranked at 1 as singletons while R. pakistanica was found dominant and ranked 5 in rank abundance (Figure 130). The median of the abundance of taxa was 5 with 2 and 13 whiskers at the lowest and highest value. The abundance of taxa in first and third quartile was 4 and 7, respectively (Figure 132). Simpsons diversity index was calculated as 0.5203. Dominance, Margalef's richness and equitability indices were calculated as 0.4797, 0.5351 and 0.7371, respectively. Additional parameters for alpha diversity analysis of the below ground taxa from all the stands are given in Table 8.

4.9.2. Beta diversity

The stands located in all three major sampling regions (Hazara division, Malakand division and Rawalpindi division), were compared to determine the beta diversity based on Jaccard's index and Bray-curtis.

For above ground beta diversity analysis, stands were compared within and between the major regions. It was noted that the species found in stand 1 and stand 2 (Hazara division) were found unique in their respective stands giving a minimum (0) value of Jaccard's index and maximum (1) Bray-Curtis. Same pattern was followed in stand 5 and stand 6 (Rawalpindi division). In stand 3 and stand 4 (Malakand division), twelve taxa including Amanita brunneopantherina, A. flavipes, Cortinarius corrosus, Gomphidius flavostipus, Inocybe alba, I. mimica, Neoboletus luridiformis, Russula amethystina, R. anthracina, R. delica, R. pakistanica and Xerocomellus rimosus were found common, while Hebeloma angustisporium, I. oblectabilis, and Suillus convexatus were found from stand 3 and A. ahmadii, A. glarea, A. swatica, Boletus himalayensis, C. longistipus, Hortiboletus rubellus, I. flavellorimosa, I. kohistanensis, I. sp. 15, Rhozopogon flavus and Russula rubecola were inhibited in stand 4 only. The values of Jaccard's index for stand 3 and 4 were 0.46 and Bray- Curtis was 0.62 showing similarity and dissimilarity between the stands, respectively. The values of similarity and dissimilarity indices between the stands in major regions based on above ground data are given in Table 9.

206

For below ground beta diversity analysis, stands were compared within and between the major regions. It has been noted that the species found in stand 1 and stand 2 (Hazara division) were only restricted to their respective localities showing Jaccard's index value 0 and Bray-Curtis 1. In Rawalpindi division, stand 5 and stand 6 also gave the same values for similarity and dissimilarity measures. Stand 3 in comparison with stand 4 (Malakand division), Inocybe sp. 1 and I. sp. 6 were shared by both the stands while 10 species were restricted to stand 3 including Clavulina cf. cinerea, C. sp. 1, I. oblectabilis, I. sp. 2, Piloderma sp. 1, Russula amethystina, R. sp. 1, Tomentella sp. 2, T. sp. 3 and Tricholoma aurantium. From stand 4, 14 species were found including Amphinema sp. 1, Clavulicium delectabile, Clavulina sp. 1, C. sp. 2, Cortinarius oulankaensis, I, sp. 3, I. sp. 4, I. sp. 5, I. sp. 13, R. anthracina, R. pakistanica, Thelephora sp. 1, Tuber sp. 1 and Wilcoxina sp. 1. The values of Jaccard's index and Bray-Curtis between these stands were 0.08 and 0.92, respectively. The values of similarity and dissimilarity indices between the stands in major regions based on below ground data are given in Table 10.

The species accumulations curves for the estimation of actual number of species in each major region were drawn. It has been shown that the number of ectomycorrhizal taxa would be increased to the saturation point with the increase in the number of samples. Maximum number of taxa has been recovered from Malakand division based on above ground and below ground sampling effort (Figures 133 & 134) followed by Hazara division (Figures 135 & 136). From Rawalpindi division, more sampling effort is required to recover more taxa (Figures 137 & 138).

4.9.3. Gamma diversity

Overall, 65 ectomycorrhizal taxa were found in association with Himalayan cedar as fruit bodies and ectomycorrhizal morphotypes from all the sampling sites. The taxa belong to 28 genera of 20 families. Above ground taxa were represented by 33 species, belonging to 19 genera and 14 families while 36 taxa belonging to 14 genera and 11 families (Figures 139 & 140). Four taxa were found from above ground as fruit bodies with their counter parts from below ground in the form of ectomycorrhizal morphotypes (Figure 141).

207

To analyze the differences among the communities in different stands in major regions from the above and below ground data, one way ANOVA was carried out. The values indicate that the communities were distinct from each other and all the stands in terms of above and below ground data did not show significant similarities. Although a few taxa were found to be similar in below ground data but the p value 0.002 indicates a high degree of dissimilarity among the stands.

It was observed that above ground ectomycorrhizal fungal communities did not show any significant response to the environmental variables including edaphic factors and rainfall patterns. Though soil saturation sulphate and available potassium and phosphorus along with some micronutrients including manganese (Mn) and Iron (Fe) have an influence on the above ground community. Below ground community structure was found dependent on edaphic factors. Most of the species were found in strong relationship with soil variables. Among these Inocybe was found cosmopolitan in distribution (Figures 142 & 143).

208

Cortinarius longistipus

Inocybe alboflavella

Taxa Inocybe alba

Geopora pinyonensis

0 5 10 15 20 25 30 Absolute abundance

Figure 98. Graph showing absolute abundance of above ground ectomycorrhizal fungal taxa in stand 1.

3% 6%

10% Geopora pinyonensis Inocybe alba Inocybe albflavella Cortinarius longistipus

81%

Figure 99. Pie chart showing relative abundance of above ground ectomycorrhizal fungal taxa in stand 1.

209

6

5

4

3

2 Rank abundance

1

0 1 2 3 Taxa

Figure 100. Rank abundance curve of above ground ectomycorrhizal fungal taxa in stand 1.

Inocybe sp. 14

Inocybe sp. 11

Trichophea sp. 1

Taxa Inocybe sp. 9

Inocybe sp. 10

Inocybe sp. 13

0 2 4 6 8 10 12 14 16 Absolute abundance

Figure 101. Graph showing absolute abundance of below ground ectomycorrhizal fungal taxa in stand 1.

210

11%

28% Inocybe sp. 13 13% Inocybe sp. 10 Inocybe sp. 9 Trichophea sp. 1 13% Inocybe sp. 11

20% Inocybe sp. 14 15%

Figure 102. Pie chart showing relative abundance of below ground ectomycorrhizal fungal taxa in stand 1.

1.5

1

0.5 Rank abundance

0 1 2 3 4 5 6 7 Taxa

Figure 103. Rank abundance curve of below ground ectomycorrhizal fungal taxa in stand 1.

211

Morchella pakistanica

Gyromitra khanspurensis Taxa

Peziza khanspurensis

Verpa asiatica

0 1 2 3 4 5 Absolute abundance

Figure 104. Graph showing absolute abundance of above ground ectomycorrhizal fungal taxa in stand 2.

10%

Verpa asiatica 20% 40% Peziza khanspurensis Gyromitra khanspurensis Morchella pakistanica

30%

Figure 105. Pie chart showing relative abundance of above ground ectomycorrhizal fungal taxa in stand 2.

212

3.5

3

2.5

2

1.5

Rank abundance 1

0.5

0 1 2 3 4 Taxa

Figure 106. Rank abundance curve of above ground ectomycorrhizal fungal taxa in stand 2.

Tomentella sp. 1 Tuber sp. 2 Russula pakistanica

Inocybe sp. 5

Cortonarius leucopus Taxa Inocybe sp. 1 Inocybe sp. 7 Peziza succosella Russula livescens

0 10 20 30 40 50 Absolute abundance

Figure 107. Graph showing absolute abundance of below ground ectomycorrhizal fungal taxa in stand 2.

213

3%

4% Russula livescens 4% 23% Peziza succosella 8% Inocybe sp. 7 Inocybe sp. 1 10% Cortonarius leucopus Inocybe sp. 5 22% Russula pakistanica 11% Tuber sp. 2 15% Tomentella sp. 1

Figure 108. Pie chart showing relative abundance of below ground ectomycorrhizal fungal taxa in stand 2.

2.5

2

1.5

1 Rank abundance 0.5

0 1 2 3 4 5 6 7 8 9 10 11 12 Taxa

Figure 109. Rank abundance curve of above ground ectomycorrhizal fungal taxa in stand 2.

214

Inocybe alba Russula pakistanica Xerocomellus rimosus Suillus convexatus Neoboletus luridiformis Amanita brunneopantherina

Gomphidius flavostipus

Cortinarius corrosus Taxa Amanita flavipes Russula amethystina Inocybe oblectabilis Inocybe mimica Hebeloma angustisporium Russula delica Russula anthracina

0 5 10 15 20 25 30 Absolute abundance

Figure 110. Graph showing absolute abundance of above ground ectomycorrhizal fungal taxa in stand 3.

2% 1% 3% Russula anthracina 3% 3% Russula delica 3% Hebeloma angustisporium 25% Inocybe mimica 4% Inocybe oblectabilis 4% Russula amethystina 4% Amanita flavipes Cortinarius corrosus 6% Gomphidius flavostipus Amanita brunneopantherina 6% 24% Neoboletus luridiformis 6% Suillus convexatus 6% Xerocomellus rimosus Russula pakistanica Inocybe alba

Figure 111. Pie chart showing relative abundance of above ground ectomycorrhizal fungal taxa in stand 3.

215

7

6

5

4

3

Rank abundance 2

1

0 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 Taxa

Figure 112. Rank abundance curve of above ground ectomycorrhizal fungal taxa in stand 3.

Tomentella sp. 2 Piloderma sp. 1 Inocybe oblectabilis Tomentella sp. 3 Inocybe sp. 6 Clavulina cf. cinerea

Taxa Inocybe sp. 1 Clavulina sp. 1 Russula amethystina Russula sp. 1 Tricholoma aurantium Inocybe sp. 2 0 10 20 30 40 50 Absolute abundance

Figure 113. Graph showing absolute abundance of below ground ectomycorrhizal fungal taxa in stand 3.

216

Inocybe sp. 2 3% 2% 1% Tricholoma aurantium 4% Russula sp. 1 4% 24% Russula amethystina 6% Clavulina sp. 1 7% Inocybe sp. 1 Clavulina cf. cinerea 8% 16% Inocybe sp. 6 Tomentella sp. 3 11% Inocybe oblectabilis 14% Piloderma sp. 1

Tomentella sp. 2 Figure 114. Pie chart chowing relative abundance of below ground ectomycorrhizal fungal taxa in stand 3.

2.5

2

1.5

1 Rank abundance 0.5

0 1 2 3 4 5 6 7 8 9 10 11 12 Taxa

Figure 115. Rank abundance curve of below ground ectomycorrhizal fungal taxa in stand 3.

217

Russula rubecola Inocybe sp. 15 Inocybe flavellorimosa Inocybe alba Hortiboletus rubellus Amanita glarea Xerocomellus rimosus Amanita ahmadii Amanita brunneopantherina Russula delica

Russula anthracina

Inocybe mimica Taxa Gomphidius flavostipus Cortinarius longistipus Boletus himalayensis Russula pakistanica Cortinarius corrosus Amanita swatica Inocybe kohistanensis Rhozopogon flavus Russula amethystina Amanita flavipes Neoboletus luridiformis

0 2 4 6 8 10 12 14 16 18 20 Absolute abundance Figure 116. Graph showing absolute abundance of above ground ectomycorrhizal fungal taxa in stand 4.

1% 1% 1% Neoboletus luridiformis 2% Amanita flavipes 1% 1% 1% Russula amethystina 3% Rhozopogon flavus 3% 16% Inocybe kohistanensis Amanita swatica 4% Cortinarius corrosus Russula pakistanica 4% Boletus himalayensis Cortinarius longistipus 4% 14% Gomphidius flavostipus 4% Inocybe mimica Russula anthracina 4% Russula delica Amanita brunneopantherina 4% Amanita ahmadii 10% 5% Xerocomellus rimosus Amanita glarea 5% 6% Hortiboletus rubellus 5% 5% Inocybe alba Inocybe alboflavella Inocybe sp. 15 Russula rubecola Figure 117. Pie chart showing relative abundance of above ground ectomycorrhizal fungal taxa in stand 4.

218

8

7

6

5

4

3 Rank abundance 2

1

0 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 Taxa

Figure 118. Rank abundance curve of above ground ectomycorrhizal fungal taxa in stand 4.

Clavulina sp. 1 Clavulina sp. 2 Russula pakistanica Wilcoxina sp. 1 Thelephora sp. 1 Amphinema sp. 1 Tuber sp. 1

Russula anthracina

Taxa Inocybe sp. 3 Inocybe sp. 13 Inocybe sp. 6 Inocybe sp. 4 Cortinarius oulankaensis Inocybe sp. 1 Inocybe sp. 5 Clavulicium delectabile

0 10 20 30 40 50 60 70 80 Absolute abundance

Figure 119. Graph showing absolute abundance of below ground ectomycorrhizal fungal taxa in stand 4.

219

Clavulicium delectabile 1% 2% 2% 3% Inocybe sp. 5 3% Inocybe sp. 1 3% 20% Cortinarius oulankaensis Inocybe sp. 4 3% Inocybe sp. 6 4% Inocybe sp. 13 6% Inocybe sp. 3 13% Russula anthracina 6% Tuber sp. 1 Amphinema sp. 1 6% Thelephora sp. 1 11% Wilcoxina sp. 1 8% 9% Russula pakistanica Clavulina sp. 2 Clavulina sp. 1 Figure 120. Pie chart showing relative abundance of below ground ectomycorrhizal fungal taxa in stand 4.

4.5 4

3.5

3 2.5 2

1.5 Rank abundance 1 0.5 0 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 Taxa

Figure 121. Rank abundance curve of below ground ectomycorrhizal fungal taxa in stand 4.

220

Tricholoma terreum

Russula pakistanica

Taxa Geastrum galiyensis

Russula delica

0 0.5 1 1.5 2 2.5 3 3.5 Absolute abundance

Figure 122. Graph showing absolute abundance of above ground ectomycorrhizal fungal taxa in stand 5.

14%

Russula delica 43% 14% Geastrum galiyensis Russula pakistanica Tricholoma terreum

29%

Figure 123. Pie chart showing relative abundance of above ground ectomycorrhizal fungal taxa in stand 5

221

2.5

2

1.5

1 Rank abundanve 0.5

0 1 2 3 4 Taxa

Figure 124. Rank abundance curve of above ground ectomycorrhizal fungal taxa in stand 5.

Inocybe sp. 6 Taxon

0 20 40 60 80 100 Absolute abundance

Figure 125. Graph showing absolute abundance of below ground ectomycorrhizal fungal taxa in stand 5.

Inocybe sp. 6

100%

Figure 126. Pie chart showing relative abundance of below ground ectomycorrhizal fungal taxon in stand 5.

222

Stand 5 1.2

1

0.8

0.6

0.4 Rank abundance 0.2

0 1 Taxon

Figure 127. Rank abundance curve of below ground ectomycorrhizal fungal taxa in stand 5.

Inocybe sp. 8

Russula pakistanica

Tuber sp. 1

0 5 10 15 20 25 30 35

Figure 128. Graph showing absolute abundance of below ground ectomycorrhizal fungal taxa in stand 6.

223

10% 21%

Tuber sp. 1 Russula pakistanica Inocybe sp. 8

69%

Figure 129. Pie chart showing relative abundance of below ground ectomycorrhizal fungal taxa in stand 6.

6

5

4

3

2 Rank abundance

1

0 1 2 3 Taxa

Figure 130. Rank abundance curve of below ground ectomycorrhizal fungal taxa in stand 6

224

12

10

8

6

Abundance 4

2

0 Stand 1 Stand 2 Stand 3 Stand 4 Stand 5 Stand 6 Stands

Figure 131. Box and whisker plot showing the number and distribution of above ground taxa in different stands.

90 80 70

60 50 40

Abundance 30 20 10 0 Stand 1 Stand 2 Stand 3 Stand 4 Stand 5 Stand 6 Stands

Figure 132. Box and whisker plot showing the number and distribution of below ground taxa in different stands.

225

Table 8. Stand wise above and belowground alpha diversity of the taxa

Buzas Berger-Parker Margalef and Stand Simpson Dominance Shannon Menhinick Gini Equitability Dominance Richness Gibson's No. Index Index Index Index Coeffificient Index Index Index Index

Above ground Stand 1 0.6538 0.3462 0.8065 0.6871 0.8736 0.7184 0.497 0.7226 0.4956 Stand 2 0.2222 0.7778 0.4 1.28 1.303 1.265 0.899 1.4 0.9232 Stand 3 0.1359 0.8641 0.25 2.287 3.014 1.471 0.6566 5.456 0.8447 Stand 4 0.07039 0.9296 0.1636 2.802 4.68 2.193 0.7167 9.262 0.8938 Stand 5 0.1905 0.8095 0.4286 1.842 1.542 1.512 0.8965 1.371 0.9212 Stand 6 ------Below ground Stand 1 0.1712 0.8288 0.2778 1.737 1.253 0.8165 0.9464 2.683 0.9692 Stand 2 0.1495 0.8505 0.2244 2.002 1.503 0.6286 0.8223 3.771 0.911 Stand 3 0.1313 0.8687 0.2396 2.185 2.092 0.866 0.7408 4.688 0.8793 Stand 4 0.09804 0.902 0.1988 2.51 2.571 0.8652 0.7694 6.583 0.9055 Stand 5 ------Stand 6 0.5203 0.4797 0.6905 0.8098 0.5351 0.4629 0.7491 0.6071 0.7371

226

Table 9. Similarity and dissimilarity indices of above ground data.

Comparative Stands Jaccard's index Bray-Curtis Stand 1 Stand 2 0.00 1.00 Stand 1 Stand 3 0.06 0.99 Stand 1 Stand 4 0.13 0.96 Stand 1 Stand 5 0.00 1.00 Stand 1 Stand 6 0.00 1.00 Stand 2 Stand 3 0.00 1.00 Stand 2 Stand 4 0.00 1.00 Stand 2 Stand 5 0.00 1.00 Stand 2 Stand 6 0.00 1.00 Stand 3 Stand 4 0.46 0.62 Stand 3 Stand 5 0.12 0.93 Stand 3 Stand 6 0.00 1.00 Stand 4 Stand 5 0.08 0.93 Stand 4 Stand 6 0.00 1.00 Stand 5 Stand 6 0.00 1.00

Table 10. Similarity and dissimilarity indices of below ground data.

Comparative Stands Jaccard's index Bray-Curtis Stand 1 Stand 2 0.00 1.00 Stand 1 Stand 3 0.00 1.00 Stand 1 Stand 4 0.05 0.92 Stand 1 Stand 5 0.00 1.00 Stand 1 Stand 6 0.00 1.00 Stand 2 Stand 3 0.05 0.93 Stand 2 Stand 4 0.14 0.83 Stand 2 Stand 5 0.00 1.00 Stand 2 Stand 6 0.09 0.93 Stand 3 Stand 4 0.08 0.92 Stand 3 Stand 5 0.08 0.94 Stand 3 Stand 6 0.00 1.00 Stand 4 Stand 5 0.06 0.95 Stand 4 Stand 6 0.12 0.83 Stand 5 Stand 6 0.00 1.00

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14

12

10 S(est) 8

6 S(est) 95% CI Lower Bound

Numberofspecies 4 S(est) 95% CI Upper 2 Bound

0 0 10 20 30 40 50 Number of individuals

Figure 133. Above ground species accumulation curve for Hazara division.

25

20

15 S(est)

10 S(est) 95% CI Lower Bound

Numberof species S(est) 95% CI Upper 5 Bound

0 0 100 200 300 Number of individuals

Figure 134. Below ground species accumulation curve for Hazara division.

228

35

30

25 S(est) 20

15 S(est) 95% CI Lower Bound

Numberofspecies 10 S(est) 95% CI Upper 5 Bound

0 0 50 100 150 200 250 Number of individuals

Figure 135. Above ground species accumulation curve for Malakand division.

35

30

25 S(est) 20

15 S(est) 95% CI Lower Bound

Numberof species 10 S(est) 95% CI Upper 5 Bound

0 0 100 200 300 400 500 Number of individuals

Figure 136. Below ground species accumulation curve for Malakand division.

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8 7

6

5 S(est) 4 S(est) 95% CI Lower 3 Bound

Numberofspecies 2 S(est) 95% CI Upper Bound 1 0 0 2 4 6 8 Number of individuals

Figure 137. Above ground species accumulation curve for Rawalpindi division.

7

6

5 S(est) 4

3 S(est) 95% CI Lower Bound

Numberofspecies 2 S(est) 95% CI Upper 1 Bound

0 0 50 100 150 Number of individuals

Figure 138. Below ground species accumulation curve for Rawalpindi division.

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60 Xerocomellus rimosus

Verpa asiatica

Tricholoma terreum

Suillus convexatus

Russula rubecola

50 Russula pakistanica Russula delica

Russula anthracina

Russula amethystina

Rhizopogon flavus

Peziza khanspurensis 40 Neoboletus luridiformis

Morchella pakistanica

Inocybe sp. 15

Inocybe oblectabillis

Inocybe mimica 30 Inocybe kohistanensis

Inocybe flavellorimosa

Inocybe alba

Hortiboletus rubellus

Hebeloma angustisporium

20 Gyromitra khanspurensis

Gomphidius flavostipus

Geopora pinyonensis

Geastrum galiyensis

Cortonarius longistipus

10 Cortinarius corrosus Boletus himalayensis

Amanita swatica

Amanita glarea

Amanita flavipes

Amanita brunneopantherina 0 Stand 1 Stand 2 Stand 3 Stand 4 Stand 5 Stand 6 Amanita ahmadii

Figure 139. Above ground community composition and comparison of stands based on frequency of taxa.

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25 Wilcoxina sp. 1 Tuber sp. 2 Tuber sp. 1 Trichophaea sp. 1 Tricholoma aurantium Tomentella sp. 3 Tomentella sp. 2 Tomentella sp. 1 20 Thelephora sp. 1 Russula sp. 1 Russula pakistanica Russula livescens Russula anthracina Russula amethystina Piloderma sp. 1 15 Peziza succosella Inocybe sp. 9 Inocybe sp. 8 Inocybe sp. 7 Inocybe sp. 6 Inocybe sp. 5 Inocybe sp. 4 10 Inocybe sp. 3 Inocybe sp. 2 Inocybe sp. 14 Inocybe sp. 12 Inocybe sp. 11 Inocybe sp. 10 Inocybe sp. 1 5 Inocybe sp 13 Inocybe obletabilis Cortinarius oulankaesis Cortinarius leucopus Clavulina sp. 2 Clavulina sp. 1 Clavulina cf. cinerea 0 Clavulicium delectabile Stand 1 Stand 2 Stand 3 Stand 4 Stand 5 Stand 6 Amphinema sp. 1 Figure 140. Below ground community composition and comparison of stands based on frequency of taxa.

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Geastrum Boletus Hortiboletus Rhizopogon Xerocomellus Suillus Neoboletus Gomphidius Hebeloma Morchella Gyromitra Verpa Geopora Above and below ground Wilcoxina Above ground Thelephora Above and below ground Clavulicium Below ground Piloderma Trichophea Amphinema Tricholoma Peziza Tuber Cortinarius Clavulina Tomentella Amanita Russula Inocybe 10 6 2 2 6 10 14 18

Figure 141. Above and below ground representative genera of ectomycorrhizal fungal taxa.

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Figure 142: Canonical correspondence analysis of above ground data with edaphic factors. Arrow ( ) showing soil variables and cross (x) showing taxa.

Figure 143. Canonical correspondence analysis of below ground data with edaphic factors. Arrow ( ) showing soil variables and cross (x) showing taxa.

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DISCUSSION

Present study represents a first glance at the ectomycorrhizal fungal communities hosted by Cedrus deodara from different regions of Pakistan. As described by Gardes & Bruns (1996), above and below ground community need to be considered simultaneously to understand the fungal communities. Fungal samples in the form of fruit bodies as well as ectomycorrhizal morphotypes were collected from three administrative divisions (Hazara, Malakand and Rawalpindi) in Khyber Pakhtunkhwa and Punjab province. From each division, two stands were selected as sampling sites. Taxa were indentified on the basis of molecular analysis as well as morphological characters. For most of the fruit bodies, molecular identification is based on ITS and LSU sequences. For some taxa where ITS was not proven to be sufficient tool, protein coding genes have also been used for the identification of taxa. ITS has been proven as a universal bar code for the identification of fungi (Schoch et al., 2012). It has been widely used for the identification of ectomycrrhizal fungi from the morphotypes. Since most of the ectomycorrhizal fungi belong to and , ITS has been successful up to 100% to amplify the DNA of these fungi from soil and ectomycorrhizae (Seifert, 2009; Porter et al., 2016)

During the field survey, 262 ectomycorrhizal fruit bodies were observed and identified. These ectomycorrhizal taxa belong to 14 families, 19 genera and 33 species. Total 919, morphotypes were isolated from the soil and 36 species belonging to 14 genera and 11families were identified. Some of the taxa were represented by the morphotypes and others as sporocarps. Only a few were found as both, above and below ground representatives.

In this study, ectomycorrhizal fungal community consisted of a few most frequent fungi and numerous rare taxa. This pattern is common for most ectomycorrhizal communities (Horton & Bruns, 2001; Smith et al., 2002; Taylor, 2002; Walker et al., 2008; Wang & Guo 2010; Wang et al., 2011). From Hazara division, samples were collected from Khanian and Khanspur; two localities geographically distant from each other. Four ectomycorrhizal fungal genera represented the ectomycorrhizal fungal community in stand 1 (Khanian). Inocybe was found to be among the most abundant, dominant and diverse genus in this area represented by seven species, two in the form of fruit bodies and five in the form of operational

235 taxonomic units (OTUs). Inocybe sp. 13 was found to be among the most abundant species in this stand covering 23.43% belowground part of the community. Geopora pinyonensis was found abundant in the form of representing 80.64% of above ground community. So, in stand 1 the community was mainly represented by Geopora and Inocybe.

In stand 2 (Khanspur), nine ectomycorrhizal fungal genera were represented by 13 species. Among these, Inocybe was found to be the most abundant, dominant and diverse genus represented by three species in the form of ectomycorrhizal morphotypes. Russula was represented by two species as morphotypes. While Peziza representing two species one (P. khanspurensis) as ascomata and other (P. succosella) as OTU. These two Peziza spp. also covered the major part of the community in terms of abundance. Cortinarius was also found from the above ground as C. longistipus and from the below ground as C. leucopus. As a whole, beta diversity at Hazara division was represented by 24 species belonging to 11 genera. Among these Inocybe was found most prevailing genus.

From Malakand division, Kalam and Mashkun were selected as sampling stands. From stand 3 (Kalam), 13 genera of ectomycorrhizal fungi were identified represented by 27 species. Inocybe was found among the major community component represented by six species followed by Russula with five species. Inocybe oblectabilis and Russula amethystina were represented from the above ground collections as well as in the form of morphotypes. Inocybe sp. 2 was found as most abundant taxon covering 46% of the below ground community. While Russula anthracina was found forming major component (25%) of the above ground in stand 1. Thus the community in stand 3 was represented by Inocybe and Russula from the above and below ground respectively.

From stand 4 (Mashkun), 37 species were identified belonging to 16 genera represented the community. Neoboletus luridiformis was found among the major above ground community component. It showed 16.36% abundance followed by Amanita flavipes with 13.63% abundance. Russula amethystina also covered a significant part of the above ground community by showing 10% abundance. Inocybe and Russula were the diverse genera at this site represented by 11 and five species respectively. Russula amethystina and R. pakistanica were represented by basidiomata as well as in the form of morphotypes. Clavulicium delectabile was among the abundant taxa covering 19.88% of the below ground

236 community while I. sp. 5, I. sp. 1 and Cortinarius oulankaensis also showed a considerable abundance with a value of 46, 39 and 30% respectively. The overall diversity at Malakand division was represented by 20 genera and 47 species. Among these, Inocybe was the major part of the community followed by Russula in the form of fruit bodies as well as morphotypes.

From Rawalpindi division, samples were collected from Kuzah Gali and Patriata. From Kuzah Gali (stand 5), four ectomycorrhizal taxa were identified belonging to four genera. Russula delica was the major component of the community with 42.85% relative abundance followed by Geastrum galiyensis with 28.57% relative abundance. From below ground, Inocybe sp. 6 was the only taxon representing the community. So, a total of five species belonging to six genera were found form stand 5. Inocybe sp. 6 and R. delica were the major community components in this stand.

From Patriata (stand 6), above ground ectomycorrhizal fungal community components were not observed. From the belowground, three species belonging to three genera were found. Russula pakistanica was the most abundant taxon representing 69.04% of the community while Tuber sp. 1 and Inocybe sp. 8 were represented by 21.42 and 9.52% relative abundance respectively. The diversity at Rawalpindi division was represented by 8 species belonging to 7 genera. Thus, the community composition in Rawalpindi division was represented by Inocybe and Russula.

In this research, only a few species (Inocybe oblectabilis, Russula amethystina, R. anthracina , R. pakistanica) were described from the root tips were found in the form of fruit bodies. These results showed that the species that do not form fruit bodies could be represented abundantly as morphotypes (Gardes & Bruns,1996; Kårén et al., 1997; Gehring et al., 1998; Jonsson et al., 1999). Moreover, the species that could not found forming ectomycorrhizal association could be found in the form of sporocarps abundantly (Nieto & Carbone, 2009). The results shown in this study also followed the same pattern. Beside this, the species that do not form conspicupus fruit bodies could not be detected during sampling particularly Thelephora and Tomentella, because of their resupinate sporocarps, these species could be missed easily (Gardes & Bruns, 1996). During this study, these two taxa were not found as sporocarps but they were the part of the community as morphotypes. These

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Thelephoroid and other resupinate taxa has been an important part of ectomycorrhizal roots in coniferous communities in Europe (Taylor & Bruns 1999; Kõljalg et al., 2000; Wurzburger et al., 2001; Horton & Bruns, 2001; Nieto & Carbone, 2009).

Previous studies on ectomycorrhizal fungal communities associated with deciduous trees also found Inocybe as a consistent genus throughout the communities (Ilyas, 2013). Horton & Bruns (2001) and Douglas et al. (2005) noted that many below ground studies have found members of the and to be dominant in coniferous ecosystems worldwide. In this study, Inocybe species were among the most diverse and dominant taxon from the above ground and below ground as well followed by Russula, which is more diverse in above ground as compared to below ground. Amanita also formed a major above ground community part in terms of diversity while Neoboletus in terms of abundance. Both these taxa were not represented by their below ground components.

Host species and age, as well as soil nutrients and organic matter content also influence the distribution patterns of ectomycorrhizal species (Alvarez et al., 1979; Dighton & Mason, 1985; Bills et al., 1986; Gehring et al., 1998). Previous studies suggest that ascomycetous fungi may not be present in old growth mixed stands. Over time these species may be reduced in areas where disturbance has been absent for some time (Cullings et al., 2000). The dominance of ascomycetous fungi in 8 years old pine stands also supports the study (Nieto & Carbone, 2009). Here in this study, from Khanspur (stand 5), mostly ascomycetous fungi dominated, and collected from near the younger hosts. Moreover, from the mixed and old forests of Kalam and Mashkun, ascomycetous taxa were not found from the above ground although a few taxa were identified from morphotypes but they were quite rare. It is clear that tree age can have impacts on ectomycorrhizal fungal communities, but that these may be more or less apparent in particular forest types, notably young plantations versus old growth (Johnson et al., 2005).

It has been noted that the areas with high nutrient availability often exhibit lower ectomycorrhizal fruit bodies, morphotype abundance, and species richness when compared to nutrient poor areas (Lilleskov et al., 2001, 2002; Peter et al., 2001; Avis et al., 2003). Research on coniferous forests where organic matter is limited has shown a link between high organic matter and declines in above and below ground ectomycorrhizal species

238 richness, colonization levels, as well as changes in community composition (Lilleskov et al., 2001, 2002; Peter et al., 2001, Avis et al., 2003). This pattern was observed in present results.

In stand 1 the amount of organic matter in the soil was 0.91% higher than the other stands. Only four genera were recovered from this area. Inocybe was found dominant and species rich genus, while Cortinarius, Geopora and Trichophea were represented by only one taxon of each. Organic matter in the soils from stand 5 and 6 was 0.89%. These stands also exhibited a lower diversity of taxa. Form stand 5 only Inocybe sp. 6 was found in abundance represented below ground community and Geastrum, Russula and Tricholoma were the above ground community representatives with only a few basidiomata.

Several studies on ectomycorrhizal fungal communities in natural environments have associated soils containing lower organic matter and lower fertility with supporting different species compositions and higher abundance, but not higher species richness (Alvarez et al., 1979; Gehring & Whitham, 1994; Gehring et al., 1998). Stand 2 bearing 0.87% organic matter content in the soil, comparatively lower than that of stand 1, 5 and 6. Species diversity and richness was found quite higher in this stand represented by 9 genera and 13 species. Stand 3 and 4 with 0.81% organic matter were the most species rich and diverse stands in the form of sporocarp as well as morphotypes. From stand 3, 27 species were found belonging to 13 genera while from stand 4, 37 species were identified belonging to 16 genera.

Besides the organic matter content as a whole, phosphorus concentration has been found most variable of the soil nutrients among sites in previous studies (Tedersoo et al., 2009). Variations in the phosphorus concentration has a positive correlation. Higher amount of available phosphorus (6.7 and 6.8) in stand 3 and 4 was found. Hyper diverse communities were recorded in these two stands. While in stand 1, 2, 4 and 5 with low diversity of taxa, 4– 5.4% of available phosphorus was found showing that phosphorus would also be considered as an important nutrient controlling the fungal communities. In some studies, ectomycorrhizal root tip abundance was negatively correlated suggesting that where soil phosphorous is more available, plants may be less dependent on ectomycorrhizae for phosphorous uptake (Kluber et al., 2012). Regional differences in the study site combining with other environmental conditions and soil chemistry could be the factors impacted these

239 results. Among other considerable mineral nutrients in soil include calcium, magnesium and sodium. These nutrients were found lower in stand 3 and 4 as compared to others showing that they could be the influential factors along with others to control the belowground community composition but they did not show a relationship with sporocarp diversity.

Studies on ectomycorrhizal colonization of Andean alder at two natural forests in northwestern Argentina showed a positive influence of higher electrical conductivity on ectomycorrhizal colonization which could be related to higher availability of mineral nutrients (Becerra et al., 2005). In present studies, stands with lower electrical conductivity showed a higher level of colonization, species richness and diversity indicated a negative relation which might be explained by the fact that other soil parameters e.g., pH, and availability of organic matter content and other mineral nutrients are interdependent along with climatic factors, host tree species in the stand and geographic factors could be the mechanism effecting the electrical conductivity and its relation to community parameters.

Most previous studies examining the effects of soil pH on ectomycorrhizal fungi suggest that acidic conditions reduce ectomycorrhizal colonization and alter community structure (Dighton & Skeffington, 1987), although some have seen no influence of acidification on community structure (Rudawska et al., 1995). Results in this study also indicated that soil pH has no remarkable influence on ectomycorrhizal fungal communities. Although it was noted that the stands with soil pH range 6.1–6.5 exhibited comparatively high diversity of ectomycorrhizal taxa. Other soil parameters remain nearly equal or did not show any significant pattern to correlate with the community structure.

Soil water availability has also been reported to influence ectomycorrhizal fungal communities (Shi et al., 2002; Cavender-Bares et al., 2009). Besides this, precipitation has been recently implicated as a driver of global patterns in ectomycorrhizal diversity (Tedersoo et al., 2012). Studies on ectomycorrhizal fungal communities in the native Scots pine woods have shown a strong correlation with precipitation and soil moisture, indicating that rain fall has a significant effect on fungal communities (Lilleskov et al., 2011). The production and composition of sporocarp communities has been observed to differ over rainfall gradients (O‟Dell et al., 2000). Several taxa with variable abundances across the rainfall gradient were identified (Jarvis et al., 2013).

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It was observed that, during the monsoon season 2012, the rain fall was recorded above than normal and ranked as 12th heaviest moon soon rainfall since 1961 (Faisal et al., 2013). During this year, 18% of the total ectomycorrhizal fungi was collected. During 2013, over all monsoon rainfall in Pakistan was observed normal to slightly above normal (Faisal, 2013). The monthly rainfall activity was close to normal in Khyber Pakhtunkhwa and Punjab province. Maximum number of fungal taxa (70%) were recorded during this year . In year 2014, the rail fall during monsoon season was recorded below 20% than normal (Faisal, 2014). Only a few (12%) ectomycorrhizal fungal taxa were recorded during this year. These results show that rainfall patterns did not affect the communities. A more than normal rain fall in year 2012 and 2013 gave different proportion of fungal taxa.

Many below ground ectomycorrhizal fungal studies encounter high variation in species frequency and abundance, which can be problematic when comparing species richness and composition between stands (Horton & Bruns, 2001). Although several dominant species were observed in each stand, but the communities still had a high number of rare species. Species richness varies greatly with sample size (Waite, 2000). The estimated level of total species richness is expected to be higher in each stand as shown by the species accumulation curves. This situation indicates that under sampling was a problem especially in dryer stands (Rawalpindi division). The use of species richness estimators addressed this issue, given that our sample size represented a part of the community and not the whole.

In this study, some members of dark septate root belonging to Ascomycota were also recorded. These endophytes were previously reported to form symbiosis with conifers of temperate forests (Trevor et al., 2001; Tedersoo et al., 2003; Menkis et al., 2005). Endophytic fungi are common in arctic-alpine habitats (Cázares 1992; Haselwandter & Read 1982; Jumpponen, 1999; Väre et al., 1992). The interactions of dark septate endophytes and their hosts are controversial, having been suggested to be pathogenic, neutral or beneficial (Fernando & Currah 1996; Haselwandter & Read 1982; O‟Dell et al., 1993; Stoyke & Currah 1993; Wang & Wilcox 1985; Wilcox & Wang, 1987). Presence of these fungi along with ectomycorrhizal symbiosis could provide a new insight for other aspects of community study.

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From this study, it has been concluded that Himalayan cedar is an excellent host for several ectomycorrhizal fungi belonging to a significant proportion of ectomycorrhizal fungal groups. These have diverse physiological tolerances enabling them to survive in many habitat types and conditions (Nieto & Carbone, 2009). Inocybe has been found cosmopolitan during this study as it has been found abundant, diverse and consistent in all the stands in the form of above and below found taxa. Besides this Russula constituted the major community component in all the regions in the form of basidiomata as well as morphotypes. Presence of both of these symbionts in nearly all the stands indicted their wide ecological amplitude. Amanita along with Boletus also constituted a major part of above ground community in Kalam and Mashkun. Many fungal taxa being the rare part of the community and 69.23% of the species recorded during this study seem undescribed previously.

These results indicate that these forests have a great potential for ectomycorrhizal fungal diversity. A through exploration of each site is needed to explore the diversity up to the saturation point. The ectomycorrhizal symbionts identified during this study could be a source of information for future studies. Rapid decline of these natural forests in the country could be restored by using these fungi as an inoculum in nursery experiments. The data generated during this study could be used for morphological comparisons. Molecular data deposited in different repositories could be used as a reference for future taxonomic and phylogenetic work.

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REFERENCES

Adamčík, S., Carteret, X., & Buyck, B. (2013). Type studies on some Russula species described by C. H. Peck. Cryptogamie Mycologie, 34, 367–391. http://dx.doi.org/10.7872/crym.v34.iss2.2013.367

Afshan, N., Khalid, A. N., & Niazi, A. R. (2004). New from Sakesar Hills. Mycopath, 1, 100–104.

Agerer, R. (1986). Studies on ectomycorrhizae. II. Introducing remarks on characterization identification. Mycotaxon, 26, 473–492.

Agerer, R. (1987–1995). Colour atlas of ectomycorrhizae. Einhorn-Verlag, Schwäbisch Gmünd.

Agerer, R. (1987–2002). Colour atlas of ectomycorrhizae. Einhorn-Verlag, Schwäbisch Gmünd.

Agerer, R. (1987–2006). Colour atlas of ectomycorrhizae. Einhorn-Verlag, Schwäbisch Gmünd.

Agerer, R. (1991). Ectomycorrhizae of imbricatus on Norway their . , 1, 21–30.

Agerer, R. (1994). Characterization of Ectomycorrhiza. In: Norris, J. R., Read, D. J., & Varma, A. K. (eds.), Techniques for the study of mycorrhiza. Methods in microbiology, 23, Academic Press London, pp. 25–73.

Agerer, R. (1999a). Never change a functionally successful principle: the evolution of (Hymenomycetes, Basidiomycota) as seen from below-ground features. Sendtnera, 6, 5–91.

Agerer, R. (1999b). Anatomical characteristics of identified ectomycorrhizas: an attempt towards a natural classification. In: Varma, A., & Hock, B. (eds.). Mycorrhiza: structure, function, molecular biology, biotechnology, Springer Verlag, Berlin, pp. 633–682.

243

Agerer, R., & Beenken, L. (1998). Geastrum fimbriatum Fr. + L. Descriptions of Ectomycorrhizae, 3, 13–18.

Agerer, R., & Iosifidou, P. (2004). Rhizomorph structure of Hymenomycetes: a possibility to test DNA-based phylogenetic hypotheses? In: Agerer, R., Piepenbring, M., & Blanz, P. (eds.). Frontiers in Basidiomycote Mycology. IHW-Verlag, Eching, pp. 249–302.

Agerer, R., & Rambold, G. (1998). DEEMY-a delta-based system characterization DEtermination of EctoMYcorrhizae. CD-ROM, version 1.1. Institute for Systematic Botany, Section Mycology, University of München.

Agerer, R., Beenken, L., & Ammirati, J. (1998). Polyporoletus sublividus Snell + Abies amabilis Forb. Descriptions of Ectomycorrhizae, 3, 85–91.

Agerer. R., Klostermeyer, D., Steglich, W., Franz, F., & Acker, G. (1996). Ectomycorrhizae of Albatrellus ovinus (Scutigeraceae) on Norway spruce with some remarks on the systematic position of the family. Mycotaxon, 59, 289–307.

Ahmad, S., Iqbal, S. H. & Khalid, A. N. (1997). Fungi of Pakistan. Sultan Ahmad Mycological Society Pakistan, pp. 1–248.

Ahmed, M., Hussain, T., Sheikh, A. H., Hussain, S. S., & Siddiqui, M. F. (2006). Phytosociology and structure of Himalayan forests from different climatic zones of Pakistan. Pakistan Journal of Botany, 38, 361–383.

Ahmed, M., Nazim, K., Siddiqui, M. F., Wahab, M., Khan, N., Khan, M.U., & Hussain, S.S. (2010). Community description of deodar forests from Himalayan range of Pakistan. Pakistan Journal of Botany, 42, 3091–3102.

Ahmed, M., Shaukat, S. S., & Siddiqui, M. F. (2011). A multivariate analysis of the vegetation of Cedrus deodara forests in Hindu Kush Himalayan ranges of Pakistan: evaluating the structure dynamics. Turkish Journal of Botany, 35, 419–438.

Allen, E. B., Allen, M. F., Helm, D. J., Trappe, J. M., Molina, R., & Rincon, E. (1995). Patterns regulation of mycorrhizal plant fungal diversity. Plant Soil, 170, 47–62.

244

Alvarez, I. F., Rowney, D. L., & Cobb Jr, F. W. (1979). Mycorrhizae and growth of white seedlings in mineral soil with and without organic layers in a California forest. Canadian Journal of Forest Research, 9, 311–315.

Ammirati, J. F., Hughes, K. W., Liimatainen, K., Niskanen, T., & Matheny, P. B. (2013). Cortinarius hesleri from eastern and related species from Europe and western North America. Botany, 91, 91–98.

Arnold, A. E., Henk, D. A., Eells, R., Lutzoni, F., & Vilgalys, R. (2007). Diversity and phylogenic affinities of foliar fungal endophytes in loblolly pine inferred by culturing and environmental PCR. Mycologia, 99, 185–206.

Arora, D., & Nguyen, N. (2014). A new species of Russula, subgenus Compactae from California. North American Fungi, 9, 1–7.

Ashraf, T., Hanif, M., & Khalid, A. N. (2012). Peziza michelii its ectomycorrhizae with Alnus nitida (Betulaceae) from Pakistan. Mycotaxon, 120, 181–188.

Avis, P. G., McLaughlin, D. J., Dentinger, B. C., & Reich, P. B. (2003). Long‐term increase in nitrogen supply alters above‐and below‐ground ectomycorrhizal communities and increases the dominance of Russula spp. in a temperate savanna. New Phytologist, 160, 239–253.

Avis, P., Muller, G., & Lussenhop, J. (2008). Ectomycorrhizal fungal communities in two North American oak forests respond to nitrogen addition. New Phytologist, 179, 472– 483.

Bahram, M., Põlme, S., Kõljalg, U., & Tedersoo, L. (2011). A single European aspen (Populus tremula) tree individual may potentially harbour dozens of Cenococcum geophilum ITS genotypes and hundreds of species of ectomycorrhizal fungi. FEMS Microbiology Ecology, 75, 313–320.

Bahram, M., Põlme, S., Kõljalg, Zerre. S., & Tedersoo, L. (2012). Regional and local patterns of ectomycorrhizal fungal diversity and community structure along an

245

altitudinal gradient in the Hyrcanian forests of northern Iran. New Phytologist, 193, 465–473.

Baquar, S. R. (1989). Medicinal Poisonous Plants of Pakistan. PRINTAS, Karachi, Pakistan, pp. 94–95.

Baseia, I. G., & Milanez, A. I. (2002). Rhizopogon (Rhizopogonaceae): hypogeous fungi in exotic plantations from the State of São Paulo, Brazil. Acta Botanica Brasilica, 16, 55–59.

Becerra, A., Pritsch, K., Arrigo, N., Palma, M., & Bartoloni, N. (2005). Ectomycorrhizal colonization of Alnus acuminata Kunth in northwestern Argentina in relation to season and soil parameters. Annals of Forest Science, 62, 325–332.

Beenken, L. (2004a). The genus Russula. Investigations on their systematics and of ectomycorrhizae. Dissertation, University of München. http://edoc.ub.uni- muenchen.de/archive/00003175/

Beenken, L. (2004b). Russula type ectomycorrhizae. Bulletin of the Mycological Society of France, 120, 293–333.

Béreau, M., Gazel, M. & Garbaye, J. (1997). The mycorrhizal symbioses of trees in the rain forest of French Guiana. Canadian Journal of Botany, 75, 711–716.

Berger, W.H., & Parker, F.L. (1970). Diversity of planktonic Foraminifera in deep sea sediments. Science, 168, 1345–1347.

Bessette, A. E., Bessette, A. R., Trudell, S. A., & Roody, W. C. (2013). Tricholomas of North America: A Mushroom Field Guide. Austin, Texas, University of Texas Press, p. 108.

Bills, G. F., Holtzman, G. I., & Miller Jr, O. K. (1986). Comparison of ectomycorrhizal- basidiomycete communities in red spruce versus northern hardwood forests of West Virginia. Canadian Journal of Botany, 64, 760–768.

246

Bisht, D., Pandey, A., & Palni, L. M. S. (2003). Influence of microbial inoculations on Cedrus deodara in relation to survival, growth promotion and nutrient uptake of seedlings and general soil microflora. Journal of Sustainable Forestry, 17, 37–54.

Bizio, E., & Ferisin, G. (2013). Inocybe mimica e altre specie rare o poco conosciute raccolte nel parco cittadino di cervignano del friuli. Bollettiino del Centro Micologico Friulano, 2013, 19–29.

Bojantchev, D., & Davis, R. M. (2013). Amanita augusta, a new species from California and the Pacific Northwest. North American Fungi, 8, 1–11.

Bon, M. (1988). Champignons d’Europe occidentale. Arthaud, Paris, p. 368.

Bon, M. (1997). Clé monographique du genre Inocybe (Fr.) Fr. Documents Mycologique, 27, 1–51.

Borovička, J., Bušek, B., Mikšík, M., Dvořák, D., Jeppesen, T. S., Dima, B., ... & Frøslev, T. G. (2015). Cortinarius prodigiosus-a new species of the subgenus Phlegmacium from Central Europe. Mycological Progress, 14, 1–8.

Boyle, H., Zimdars, B., Renker, C., & Buscot, F. (2006). A molecular phylogeny of Hebeloma species from Europe. Mycological Research, 110, 369–380.

Brand, F. (1992). Mixed associations of fungi in ectomycorrhizal roots. In: Read, D.J., Lewis, D.H., Fitter, A. H., & Alexer, I. J. (eds.), Mycorrhizas in Ecosystems, CAB International, Wallingford, UK, pp. 142–147.

Bray, J. R., & Curtis J. T. (1957). An ordination of upland forest communities of southern Wisconsin. Ecological Monographs, 27, 325–349.

Breitenbach, J., & Kränzlin, F. (2000). Fungi of Switzerland (vol. 5 Lucerne edn.). Mykologia Lucerne.

Britzelmayr, M. (1890). Hymenomycetes from Südbayern 6 : Boleti , Cortinarii , Dermini , Hydnei , Hyporhodii , Leusospori , Melanospori . Reports of the Natural Sciences Association Swabia Neuburg, 30, 1–34.

247

Brundrett, M. C. (2009). Mycorrhizal associations other means of nutrition of vascular plants: understanding the global diversity of host plants by resolving conflicting information developing reliable means of diagnosis. Plant Soil, 320, 37–77.

Bruns, T. D. (1995). Thoughts on the processes that maintain local species diversity of ectomycorrhizal fungi. In The significance and regulation of soil biodiversity. Springer Netherlands, pp. 63–73.

Bruns, T. D., White, T. J., & Taylor, J. W. (1991). Fungal molecular systematics. The Annual Review of Ecology, Evolution Systematics, 22, 525–564.

Buchheim, M. A., Keller, A., Koetschan, C., Förster, F., Merget, B., & Wolf, M. (2011). Internal transcribed spacer 2 (nu ITS2 rRNA) sequence-structure phylogenetics: towards an automated reconstruction of the green algal tree of life. PloS One, 6, e16931.

Buscot, F. (1989). Field observations on growth and development of Morchella rotunda and Mitrophora semilibera in relation to forest soil temperature. Canadian Journal of Botany, 67, 589–593. http://dx.doi.org/10.1139/b89-080

Buscot, F., & Roux, J. (1987). Association between living roots and ascocarps of Morchella rotunda (Pers.) Boudier. Transactions of the British Mycological Society, 89, 249– 252. http://dx.doi.org/10.1016/S0007-1536(87)80162-6

Buscot, G. F., & Bernillion, J. (1991). Mycosporins and related compounds in field and cultural mycelial structures of Morchella esculenta. Mycological Research, 95, 752– 754. http://dx.doi.org/10.1016/S0953-7562(09)80826-5

Caboň, M., Adamčík, S., & Valachovič, M. (2013). Diversity of the family Russulaceae in the Scots pine forests of Záhorská nížina (SW Slovakia). Czech Mycology, 65, 179– 191.

Cafaro, M. J. (2005). Eccrinales (Trichomycetes) are not fungi, but a clade of protists at the early divergence of animals and fungi. Molecular Phylogenetics and Evolution, 35, 21–34.

248

Cai, Q., Tulloss, R. E., Tang, L. P., Tolgor, B., Zhang, P., Chen, Z. H., & Yang, Z. L. (2014). Multi-locus phylogeny of lethal : Implications for species diversity and historical biogeography. BMC evolutionary biology, 14, 143–158.

Cavender-Bares, J., Izzo, A., Robinson, R., & Lovelock, C. E. (2009). Changes in ectomycorrhizal community structure on two containerized oak hosts across an experimental hydrologic gradient. Mycorrhiza, 19, 133–142.

Cázares, E. (1992). Mycorrhizal fungi, their relationship to plant succession in subalpine habitats. PhD thesis, Oregon State University, Corvallis.

Champion, G. Harry, S., & Seth, K. (1965). Forest types of Pakistan. Pakistan Forest Institute, Peshawar, p. 233.

Champion, H. G., Seth. S. K., & Khattak, G. M. (1968). Forests Types of Pakistan, Pakistan Forest Institute, Peshawar, Pakistan, pp. 1–100.

Chilvers, G. A. (1968). Some distinctive types of eucalypt mycorrhiza. Australian Journal of Botany,16, 49–70.

Cline, E. T., Ammirati, J. F., & Edmonds, R. L. (2005). Does proximity to mature trees influence ectomycorrhizal fungus communities of Douglas‐fir seedlings?. New Phytologist, 166, 993–1009.

Colpaert, J. V. (2008). Heavy metal pollution and genetic adaptations in ectomycorrhizal fungi. In British Mycological Society Symposia Series, 27, Academic Press, pp. 157– 173.

Colwell, R. K., Mao, C. X., & Chang, J. (2004). Interpolating, extrapolating, and comparing incidence-based species accumulation curves. Ecology, 85, 2717–2727.

Comini, O., & Pacioni, G. (1997). Mycorrhizae of Asian black , Tuber himalayense T. indicum. Mycotaxon, 63, 77–86.

Cullings, K. W. (1992). Design and testing of a plant‐specific PCR primer for ecological and evolutionary studies. Molecular Ecology, 1, 233–240.

249

Cullings, K. W., Vogler, D. R., Parker, V. T., & Finley, S. K. (2000). Ectomycorrhizal specificity patterns in a mixed Pinus contorta and Picea engelmannii forest in yellowstone National Park. Applied and Environmental Microbiology, 66, 4988– 4991.

Dahlberg, A. (2001). Community ecology of ectomycorrhizal fungi: an advancing interdisciplinary field. New Phytologist, 150, 555–562.

Dahlstrom, J. L., Smith, J. E., & Weber, N. S. (2000). Mycorrhiza-like interaction by Morchella with species of the Pinaceae in pure culture synthesis. Mycorrhiza, 9, 279– 285. http://dx.doi.org/10.1007/PL00009992

Das, K., Atri, N. S., & Buyck B. (2013). Three new species of Russula () from India. Mycosphere, 4, 722–732. http://dx.doi.org/10.5943/mycosphere/4/4/9

Das, K., Sharma, J. R., & Atri., N. S. (2006). Russula in Himalaya 3: a new species of subgenus Ingratula. Mycotaxon, 95, 271–275.

Deepika, K., Ramesh, C. Upadhyay, M., & Reddy, S. (2011). Cantharellus pseudoformosus, a new species associated with Cedrus deodara from India. Mycoscience, 52, 147– 151.

Dentinger, B., Gaya, E., O'Brien, H., Suz, L. M., Lachlan, R., Díaz‐Valderrama, J. R., ... & Aime, M. C. (2016). Tales from the crypt: genome mining from fungarium specimens improves resolution of the mushroom tree of life. Biological Journal of the Linnean Society, 117, 11–32. de-Roman, M., Claveria, V., & de-Miguel, A. M. (2005). A revision of the descriptions of ectomycorrhizas published since 1961. Mycological research, 109, 1063–1104.

Dighton, J., & Mason, P. A. (1985). Mycorrhizal dynamics during forest tree development. Developmental biology of higher fungi. Cambridge University Press, pp. 117–139.

Dighton, J., & Skeffington, R. A. (1987). Effects of artificial acid precipitation on the mycorrhizas of Scots pine seedlings. New Phytologist, 107, 191–202.

250

Digrak, M., Alma, M. H., Iicim, A., & Sen, S. (1999). Antibacterial antifungal effect of various commercial plant extracts. Pharmaceutical Biology, 37, 216–220.

Dimri, V., & Sharma, M. C. (2004). Effects of sarcoptic mange its control with oil of Cedrus deodara, Pangamia glabra, Jatropha curcas and benzyl benzoate, both with without ascorbic acid on growing sheep, epidermiology, assessment of clinical, haematological, cell mediated himoral immune responses pathology. Journal of Veterinary Medicine Series A, 51, 71–78.

Dominik, T. (1969). Key to ectotrophic mycorrhizae. Folia Forestalia Polonica Series A, 15, 309–328.

Douglas, R. B., Parker, V. T., & Cullings, K. W. (2005). Belowground ectomycorrhizal community structure of mature lodgepole pine and mixed conifer stands in Yellowstone National Park. Forest Ecology and Management, 208, 303–317.

Durall, D. M., Jones, M. D., Wright, E. F., Kroeger, P., & Coates, K. D. (1999). Species richness of ectomycorrhizal fungi in cut blocks of different sizes in the Interior Cedar-Hemlock forests of northwestern British Columbia: sporocarps and ectomycorrhizae. Canadian Journal of Forest Research, 29, 1322–1332.

Eberhardt, U. (2002). Molecular kinship analyses of the agaricoid Russulaceae: correspondence with mycorrhizal anatomy sporocarp features in the genus Russula. Mycological Progress, 1, 201–223.

Eberhardt, U., Oberwinkler, F., Verbeken, A., Rinaldi, A. C., Pacioni, G., & Comini, O. (2000). Lactarius ectomycorrhizae on : morphological description, molecular characterization, taxonomic remarks. Mycologia, 92, 860–873.

Eberhardt, U., Walter, L., & Kottke, I. (1999). Molecular morphological discrimination between Tylospora fibrillosa and Tylospora asterophora mycorrhizae. Canadian Journal of Botany, 77, 11–21.

Egger, K. N. (1995). Molecular analysis of ectomycorrhizal fungal communities. Canadian Journal of Botany, 73, 1415–1422.

251

Egger, K. N., & Hibbett, D. S. (2004). The evolutionary implications of reciprocal exploitation in mycorrhizas. Canadian Journal of Botany, 82, 1110–1121.

Egli, S., Amiet, R., Zollinger, M., & Schneider, B. (1993). Characterization of (L.) Karst. ectomycorrhizas: discrepancy between classification according to macroscopic versus microscopic features. Trees, 7, 123–129.

Farjon, A. (1990). Pinaceae. Drawings Descriptions of the Genera. Koeltz Scientific Books.

Favre, J. (1955). Les champignons superieurs de la zone alpine du Parc National suisse. Ergebn. Wiss. Untersuch. Schweiz. National parkes 5, 1–212.

Fernando, A. A., & Currah, R. S. (1996). A comparative study of the effects of the root endophytes Leptodontidium orchidicola and Phialocephala fortinii (Fungi Imperfecti) on the growth of some subalpine plants in culture. Canadian Journal of Botany, 74, 1071–1078.

Finlay, R. D., Lindahl, B. D., & Taylor, A. F. (2008). Responses of mycorrhizal fungi to stress. In British Mycological Society Symposia Series, 27, Academic Press, pp. 201– 219.

Fisher, R. A. (1918). The Correlation Between Relatives on the Supposition of Mendelian Inheritance. Philosophical Transactions of the Royal Society of Edinburgh, 52, 399– 433.

Flores-Rentería, L., Lau, M. K., Lamit, L. J., & Gehring, C. A. (2014). An elusive ectomycorrhizal fungus reveals itself: a new species of Geopora (Pyronemataceae) associated with Pinus edulis. Mycologia, 106, 553–563.

Frank, A. B. (1885). On the nutrient providing root-symbiosis between underground fungi and certain trees. Journal of the American Botanical Society, 3, 128–145.

Fries, E.M. (1822). Systema Mycologicum II. 1, 61.

Fries, E.M. (1838). Epicrisis Systematis Mycologici. Synopsis Hymenomycetum, i–xii, 1– 612.

252

Gamito, S. (2010). Caution is needed when applying Margalef diversity index. Ecological Indicators, 10, 550–551.

Gardes, M., & Bruns, T. D. (1993). ITS primers with enhanced specificity for basidiomycetes application to the identification of mycorrhizae rusts. Molecular Ecology, 2, 113–118.

Gardes, M., & Bruns, T. D. (1996). Community structure of ectomycorrhizal fungi in a Pinus muricata forest: above belowground views. Canadian Journal of Botany, 74, 1572– 1583.

Ge, C., Cui, Y. N., Jing, P. Y., & Hong, X. Y. (2014). An alternative suite of universal primers for genotyping in multiplex PCR. PloS One, 9, e92826.

Ge, Z. W., Smith, M. E., Zhang, Q. Y., & Yang, Z. L. (2012). Two species of the Asian endemic genus Keteleeria form ectomycorrhizas with diverse fungal symbionts in southwestern China. Mycorrhiza, 22, 403–408.

Ge, Z. W., Yang, Z. L., & Vellinga, E. C. (2010). The genus Macrolepiota (, Basidiomycota) in China. Fungal Diversity, 45, 81–98.

Ge, Z. W., Yang, Z. L., Qasim, T., Nawaz, R., Khalid, A. N., & Vellinga, E. C. (2015). Four new species in Leucoagaricus (Agaricaceae, Basidiomycota) from Asia. Mycologia, 107, 1033–1044.

Gehring, C. A., & Whitham, T. G. (1994). Comparisons of ectomycorrhizae on pinyon (Pinus edulis; Pinaceae) across extremes of soil type and herbivory. American Journal of Botany, 81, 1509–1516.

Gehring, C. A., Theimer, T. C., Whitham, T. G., & Keim, P. (1998). Ectomycorrhizal fungal community structure of pinyon pines growing in two environmental extremes. Ecology, 79, 1562–1572.

Gelardi, M., Simonini, G., & Vizzini, A. (2014). Nomenclatural novelties. Index Fungorum, 192.

253

Geml, J., Laursen, G. A., Timling, I., McFarland, J. M., Booth, M. G., Lennon, N., ... & Taylor, D. (2009). Molecular phylogenetic biodiversity assessment of arctic and boreal ectomycorrhizal Lactarius Pers. (Russulales; Basidiomycota) in Alaska, based on soil and sporocarp DNA. Molecular Ecology, 18, 2213–2227.

Gibelli, G. (1883). New studies on the disease called Castagno detta dell‟ inchiostro. Memories of the Academy of Sciences Institute of Bologna, 4, 287–314.

Gini, C. (1912). Variability and Mutability. Journal of the Royal Statistical Society, 76, 326– 327.

Goodman, D. M., Durall. D. M., Trofymow, J. A., & Berch, S. (1996–2000). A manual of Concise Description of North American Ectomycorrhizae: including microscopic molecular characterization. Mycologue Publications, Sydney, pp. 38–99.

Gottlieb, A. M., & Lichtwardt, R. W. (2001). Molecular variation within and among species of Harpellales. Mycologia, 93, 66–81.

Grey, P. (2005). Fungi down under: the fungimap guide to Australian fungi. Fungimap section. Royal Botanic Gardens Melbourne, pp. 1–146.

Gronbach, E. (1988). Characterization and identification of ectomycorrhizae in a Fichtenbest with studies of trait variability in acid-irrigated areas. Bibliotheca Mycologica, 125, 1–217.

Gupta, S., Walia, A., & Malan, R. (2010). Phytochemistry and pharmacology of Cedrus deodera: An overview. International Journal of Pharmaceutical Sciences and Research, 2, 2010–2020.

Haas, H. (1969). The young specialist looks at Fungi. Burke, p. 74.

Hall, I. R., Yun, W., & Amicucci, A. (2003). Cultivation of edible ectomycorrhizal . Trends in Biotechnology, 21, 433–438.

Hall, T. A. (1999). BioEdit: a user-friendly biological sequence alignment editor and analysis program for Windows 95/98/NT. Nucleic acids symposium series, 4, 95–98.

254

Hamayun, M., Khan, M. A, & Begum, S. (2003). Marketing of medicinal plants of Utror- Gabral valleys, Swat, Pakistan. Ethnobotanical Leaflet 2003, 13.

Hanif, M. (2012). Aphyllophorales and their morphotypes from Pakistan. PhD. Thesis, Department of Botany, University of Punjab, Lahore, Pakistan

Hanif, M., Khalid, A. N., & Sarwar, S. (2012). Additions to the ectomycorrhizae associated with Himalayan cedar (Cedrus deodara) using rDNA-ITS. International Journal of Agriculture Biology, 13, 1062–1067.

Hansen, K., Læssøe, T., & Pfister, D. H. (2002). Phylogenetic diversity in the core group of Peziza inferred from ITS sequences and morphology. Mycological Research, 106, 879–902.

Harmaja, H. (1973). Amendments of the limits of the genera Gyromitra and Pseudorhizina, with the description of a new species, Gyromitra montana. Karstenia, 13, 48–58.

Harrower, E., Bougher, N. L., Winterbottom, C., Henkel, T. W., Horak, E. & Matheny, P. B. (2015). New species in Cortinarius section Cortinarius () from the Americas and Australasia. MycoKeys, 11, 1.

Haselwandter, K., & Read, D. J. (1982). The significance of a root-fungus association in two Carex species of high-alpine plant communities. Oecologia, 53, 352–354.

Haug, I., Weiß, M., Homeier, J., Oberwinkler, F., & Kottke, I. (2004). Russulaceae Thelephoraceae form ectomycorrhizas with members of the Nyctaginaceae (Caryophyllales) in the tropical mountain rain forest of southern Ecuador. New Phytologist, 165, 923–936.

Hayek, L. A., & Buzas, M. A. (2010). Surveying natural populations. Columbia University Press, pp. 1–590.

Healy, R. A., Huffman, D. R., Tiffany, L. H., & Knaphaus, G. (2008). Mushrooms and other fungi of the Midcontinental United States (Bur Oak Guide). Iowa City, Iowa, University of Iowa Press. p. 295.

255

Heim, R. (1931). Le genre inocybe: précédé d'une introduction générale à l'étude des ochrosporés. Lechevalier, p. 431.

Heluta, V. P. (2001). Gyromitra slonevskii Heluta – A new discomycete (Pezizales) from Ukraine. Ukrainian Botanical Journal, 58, 83–85.

Henkel, T. W., Terborght, J., & Vilgalys, R. (2002). Ectomycorrhizal fungi their leguminous hosts in the Pakaraima Mountains of Guyana. Mycological Research, 106, 515–531.

Hesler, L. R. (1977). New species of Hebeloma. Kew Bulletin, 31, 471–480.

Hibbett, D. S., & Matheny, P. B. (2009). The relative ages of ectomycorrhizal mushrooms their plant hosts estimated using Bayesian relaxed molecular clock analyses. BMC Biology, 7, 13.

Hibbett, D. S., Pine, E. M., Langer, G., Langer, E., & Donoghue, M. J. (1997). Evolution of gilled mushrooms and puffballs inferred from ribosomal DNA sequences. Proceedings of the National Academy of Sciences, 94, 12002–12006.

Horton, T. R., & Bruns, T. D. (2001). The molecular revolution in ectomycorrhizal ecology: peeking into the black-box. Molecular Ecology, 10, 1855–1871.

Horton, T. R., Molina, R., & Hood, K. (2005). Douglas-fir ectomycorrhizae in 40-and 400- year-old stands: mycobiont availability to late successional western hemlock. Mycorrhiza, 15, 393–403.

http://www.mushroomexpert.com/gyromitra_montana.html

Hussain, F., & Illahi, I. (1991). Ecology Vegetation of Lesser Himalayan Pakistan. Botany Department, University of Peshawar, pp. 187.

Ilyas, S. (2013). Molecular investigations to characterize ectomycorrhizal fungal communities associated with some deciduous trees of Galyat, Pakistan. PhD. Thesis, Department of Botany, University of Punjab, Lahore, Pakistan, pp. 1–150.

256

Ingleby, K., Mason, P. A., Last, F. T., & Fleming, L. V. (1990). Identification of Ectomycorrhizas. Institute of Terrestrial Ecology. Research Publication No. 5. HMSO, London, pp. 69–141.

Izzo, A., Agbowo, J., & Bruns, T. D. (2005). Detection of plot‐level changes in ectomycorrhizal communities across years in an old‐growth mixed‐conifer forest. New Phytologist, 166, 619–630.

Jabeen, S. Ashraf, T., & Khalid, A. N. (2015a). Peziza succosella and its ectomycorrhiza associated with Cedrus deodara from Himalayan moist temperate forests of Pakistan. Mycotaxon, 130, 455–464.

Jabeen, S. Sarwar, S. Niazi, A. R. & Khalid, A. N. (2014). Checklist of ectomycorrhizae from Pakistan. Annals of Applied Bio-Sciences. 1, 10–20. Jabeen, S., & Khalid, A. N. (2014). Community structure of ectomycorrhizae associated with Salix spp. growing in two different climatic regions of Pakistan. International Journal of Agriculture and Biology, 16, 480–488.

Jabeen, S., Ahmad, I., Rashid, A., & Khalid, A. N. (2016b). Inocybe kohistanensis, a new species from Pakistan. Turkish Journal of Botany, 40, 312–318.

Jabeen, S., Fiaz, M., Saba, M. Ahmad, H., & Khalid, A. N. (2015b). Molecular phylogeny and morphological characterization of Russula livescens and its ectomycorrhiza from mixed coniferous forests of Pakistan. Austrian Journal of Mycology, 23, 145–154.

Jabeen, S., Ilyas, S., Niazi, A. R., & Khalid, A. N. (2012). Diversity of ectomycorrhizae associated with Populus spp., growing in two different ecological zones of Pakistan. International Journal of Agriculture and Biology, 14, 681–688.

Jabeen, S., Niazi, A. R., & Khalid, A. N. (2016a). First record of Russula anthracina and its ectomycorrhiza associated with Himalayan cedar from South Asia. Mycotaxon, 131, 31–44.

257

Jaccard, P. (1901). Distribution de la flore alpine dans le bassin des Dranses et dans quelques régions voisines. Bulletin de la Société Vaudoise des Sciences Naturelles, 37, 241– 272.

Jarvis, S., Woodward, S., Alexander, I. J., & Taylor, A. F. S. (2013). Regional scale gradients of climate and nitrogen deposition drive variation in ectomycorrhizal fungal communities associated with native Scots pine. Global Change Biology, 19, 1688– 1696.

Jenkins, D. T. (1986). Amanita of North America. Mad River Press, Eureka, CA, pp. 1–197.

Jensen, A. B., Gargas, A., Eilenberg, J., & Rosendahl, S. (1998). Relationships of the insect- pathogenic order Entomophthorales (Zygomycota, Fungi) based on phylogenetic analyses of nuclear small subunit ribosomal DNA sequences (SSU rDNA). Fungal Genetics and Biology, 24, 325–334.

Johnson, D., Ijdo, M., Genney, D. R., Erson, I. C., & Alexer, I. J. (2005). How do plants regulate the function, community structure, diversity of mycorrhizal fungi. Journal of Experimental Botany, 56, 1751–1760.

Jonsson, L., Dahlberg, A., Nilsson, M. C., Zackrisson, O., & Kårén, O. (1999). Ectomycorrhizal fungal communities in late-successional Swedish boreal forests, their composition following wildfire. Molecular Ecology, 8, 205–215.

Judd, W. S., Campbell, C. S., Kellogg, E. A., Stevens, P. F., & Donoghue, M. J. (2002). Plant Systematics: A Phylogenetic Approach, Second Edition. Sinauer Associates, Inc., Sunderland, pp. 1–39.

Jumpponen, A. (1999). Spatial distribution of discrete RAPD phenotypes of a root endophytic fungus, Phialocephala fortinii, at a primary successional site on a glacier forefront. New Phytologist, 141, 333–344.

Kårén, O., & Nylund, J. E. (1996). Effects of N-free fertilization on ectomycorrhiza community structure in Norway spruce in southern Sweden. Plant Soil, 181, 295– 305.

258

Kårén, O., Hogberg, N., Dahlberg, A., Jonsson, L., & Nylund, J. E. (1997). Inter- and intraspecific variation in the ITS region of rDNA of ectomyocorrhizal fungi in Fennoscandia as detected by endonuclease analysis. New Phytologist, 136, 313–325.

Kårén, O., Högberg, N., Dahlberg, A., Jonsson, L., & Nylund, J. E. (2008). Inter- and intraspecific variation in the ITS region of rDNA of ectomycorrhizal fungi in Fennoscandia as detected by endonuclease analysis. New Phytologist, 136, 313–325.

Kauffman, C. H. (1924). Inocybe. North American Flora, 10, 227–261.

Kazmi, S. A. R., Khalid, A. N., & Niazi, A. R. (2004). Ectomycorrhizal diversity with Himalayan Poplar (Populus ciliata Wall ex Royle). Mycopath, 2, 75–78.

Keane, P. J., Kile, G. A., & Podger, F. D. (2000). Diseases and Pathogens of Eucalypts. Canberra: CSIRO Publishing. p. 84.

Khalid, A. N., & Niazi, A. R. (2003). New ectomycorrhizas in association with Poplar from Himalayan moist forests of Pakistan. Mycopath, 1, 95–98.

Kim, C. S., Jo, J. W., Kwag, Y. N., Kim, J. H., Shrestha, B., Sung, G. H., & Han, S. K. (2013). Taxonomic study of Amanita subgenus Lepidella and three unrecorded Amanita species in Korea. Mycobiology, 41, 183–190.

Kirk, P. M., Ainsworth, G. C., Cannon, P. F., & Minter, D. W. (2008). Ainsworth Bisby's Dictionary of Fungi (10th ed.). CAB International, UK.

Kluber, L. A., Carrino-Kyker, S. R., Coyle, K. P., DeForest, J. L., Hewins, C. R., Shaw, A. N., ... & Burke, D. J. (2012). Mycorrhizal response to experimental pH and P manipulation in acidic hardwood forests. PloS One, 7, e48946.

Kobayashi, T. (2003). Notes on the genus Inocybe of Japan: II. Mycoscience, 44, 383–388.

Kõljalg, U., Dahlberg, A., Taylor, A. F. S., Larsson, E., Hallenberg, N., Stenlid, J., ... & Jonsson, L. (2000). Diversity and abundance of resupinate thelephoroid fungi as ectomycorrhizal symbionts in Swedish boreal forests. Molecular Ecology, 9, 1985– 1996.

259

Kropp, B. R., Matheny, P. B., & Hutchison, L. J. (2013). Inocybe section Rimosae in Utah: phylogenetic affinities and new species. Mycologia, 105, 728–747.

Kühner, R. (1933). Notes sur le genre Inocybe1. Les Inocybes goinosporés (Fin). Bulletin de la Société mycologique de France, 49, 81–121.

Kühner, R. (1988). Diagnoses de quelques nouveaux Inocybes récoltés en zone alpine de la Vanoise (Alpes françaises). Documents Mycologiques, 19, 1–27.

Kumari, D., Upadhyay, R. C., & Reddy, M. S. (2012). Craterellus indicus sp. nov., a new species associated with Cedrus deodara from the western Himalayas, India. Mycological Progress, 11, 769–774.

Kuo, M. (2005). Amanita pantherina. Retrieved from the mushroomexpert.com web site : http://www.mushroomexpert.com/amanita_pantherina.html

Kuo, M. (2011). The genus Cortinarius. Retrieved from the mushroomexpert.com web site: http://www.mushroomexpert.com/cortinarius.html

Kuo, M. (2012). Gyromitra montana (Gyromitra gigas). Retrieved from the mushroomexpert.com Web site: Kuo, M. (2013). Amanita rubescens. Retrieved from the mushroomexpert.com web site: http://www.mushroomexpert.com/amanita_rubescens.html

Kuo, M. (2014). The genus Gomphidius. Retrieved from the mushroomexpert.com web site: http://www.mushroomexpert.com/gomphidius.html

Laessoe, T. (1998). Mushrooms (flexi bound). Dorling Kindersley.

Lamaison, J. L., & Polese, J. M. (2005). The great encyclopedia of mushrooms. Könemann. p. 25.

Larsson, E., Ryberg, M., Moreau, P. A., Mathiesen, Å. D., & Jacobsson, S. (2009). Taxonomy and evolutionary relationships within species of section Rimosae (Inocybe) based on ITS, LSU and mtSSU sequence data. Persoonia-Molecular Phylogeny and Evolution of Fungi, 23, 86–98.

260

Latha, K. D., & Manimohan, P. (2016). Five new species of Inocybe (Agaricales) from tropical India. Mycologia, 108, 110–122

Li, G. J. (2014). Taxonomy of Russula from China. PhD. dissertation. Institute of Microbiology, Chinese Academy of Sciences & University of Chinese Academy of Sciences (In Chinese).

Li, G. J., Zhao, D., Li, S. F., & Wen, H. A. (2015). Russula chiui and R. pseudopectinatoides, two new species from south western China supported by morphological and molecular evidence. Mycological Progress, 14, 1–14.

Liddell, H. G., & Scott, R. (1980). A Greek-English Lexicon. United Kingdom: Oxford University Press.

Lilleskov, E. A., Fahey, T. J., & Lovett, G. M. (2001). Ectomycorrhizal fungal aboveground community change over an atmospheric nitrogen deposition gradient. Ecological Applications, 11, 397–410.

Lilleskov, E. A., Fahey, T. J., Horton, T. R., & Lovett, G. M. (2002). Belowground ectomycorrhizal fungal community change over a nitrogen deposition gradient in Alaska. Ecology, 83, 104–115.

Lilleskov, E. A., Hobbie, E. A., & Horton, T. R. (2011). Conservation of ectomycorrhizal fungi: exploring the linkages between functional and taxonomic responses to anthropogenic N deposition. Fungal Ecology, 4, 174–183.

Long, D., Liu, J., Han, Q., Wang, X., & Huang, J. (2016). Ectomycorrhizal fungal communities associated with Populus simonii and Pinus tabuliformis in the hilly- gully region of the Loess Plateau, China. Scientific reports, 6.

Magurran, A. E. (2013). Measuring biological diversity. John Wiley & Sons, pp. 1–264.

Manimohan, P., & Latha, K. P. D. (2011). Observation on two rarely collected species of Russula. Mycotaxon, 116, 125–131.

261

Margalef, R., 1958. Temporal succession and spatial heterogeneity in phytoplankton. In: Perspectives in marine biology, Buzzati-Traverso, A. A. (ed.), University of California Press, Berkeley, pp. 323–347.

Massee, G. E. (1904). A monograph of the genus Inocybe, Karsten. Annals of Botany, 18, 460–502.

Masui, K. (1926). A study of the mycorrhiza of Abies firma, S. et Z., with special reference to its mycorrhizal fungus, Cantharellus floccosus, Schw. Memoirs of the College of Science; Kyoto Imperial University. Series B Biology Kyoto, 2, 15–84.

Maurer, B. A., & McGill, B. J. (2011). Biological diversity. Oxford University Press, Oxford, pp. 55–65.

May, T. W. (2003). The status of names and records of Australian macrofungi. New Zealand Journal of Botany, 41, 379–389.

Melin, E. (1923). Experimental investigations on the constitution and ecology of the mycorrhiza of Pinus silvestris L. and Picea Abies (L.) Karst. Mykologische Untersuchungen Berlin, 2, 73–331.

Menhinick, E. F. (1964). A comparison of some species individual diversity indices applied to samples of field insects. Ecology, 45, 839–861.

Menkis, A., Vasiliauskas, R., Taylor, A. F., Stenlid, J., & Finlay, R. (2005). Fungal communities in mycorrhizal roots of conifer seedlings in forest nurseries under different cultivation systems, assessed by morphotyping, direct sequencing and mycelial isolation. Mycorrhiza, 16, 33–41.

Molina, R., Massicotte, H., & Trappe, J. M. (1992). Specificity phenomena in mycorrhizal symbiosis: community-ecological consequences practical implications. In: Allen, M. F. (ed.). Mycorrhizal functioning: An integrative plant-fungal process. Chapman Hall, New York, pp. 357–423.

262

Moncalvo, J. M., Lutzoni, F. M., Rehner, S. A., Johnson, J., & Vilgalys, R. (2000). Phylogenetic relationships of agaric fungi based on nuclear large subunit ribosomal DNA sequences. Systematic Biology, 49, 278–305.

Moncalvo, J. M., Vilgalys, R., Redhead, S. A., Johnson, J. E., James, T. Y., Aime, M. C., ... & Thorn, R. G. (2002). One hundred and seventeen clades of euagarics. Molecular Phylogenetics and Evolution, 23, 357–400.

Moreau, P. A., Bellanger, J. M., Clowez, P., Courtecuisse, R., Hansen, K., Knudsen, H., ... & Richard, F. (2014). (2289) Proposal to conserve the name Morchella semilibera against Phallus crassipes, P. gigas and P. undosus (Ascomycota). Taxon, 63, 677– 678.

Morris, M. H., Smith, M. E., Rizzo, D. M., Rejmánek, M., & Bledsoe, C. S. (2008). Contrasting ectomycorrhizal fungal communities on the roots of co‐occurring (Quercus spp.) in a California woodland. New Phytologist, 178, 167–176.

Moyersoen, B., Becker, P., & Alexer, I. J. (2001). Are ectomycorrhizas more abundant than arbuscular mycorrhizas in tropical heath forests? New Phytologist, 150, 591–599.

Munsell, A. H. (1975). Munsell soil color charts. Baltimore, MD, USA.

Nataranjan K., Senthilarasu G., Kumaresan V., & Rivière T. (2005). Diversity in ectomycorrhizal fungi of a dipterocarp forest in Western Ghats. Current Science, 88, 1893–1895.

Nezzar-Hocine, H., Perrin, R., Halli-Hargas, R., & Chevalier, G. (1998). Ectomycorrhizal associations with Cedrus atlantica (Endl) Manetti ex Carrière. I. Mycorrhizal synthesis with Tricholoma tridentinum Singer var. cedretorum Bon. Mycorrhiza, 8, 47–51.

Niazi A. R., Iqbal S. H., & Khalid A. N. (2009). Ectomycorrhizae between Amanita rubescens Himalayan Spruce (Picea smithiana) from Pakistan. Mycotaxon, 107, 73– 80.

263

Niazi, A. R. (2008). Biodiversity of ectomycorrhizas in Conifers from Himalayan Moist Temperate Forests of Pakistan. PhD. Thesis, Department of Botany, University of Punjab, Lahore, Pakistan, pp. 1–248.

Niazi, A. R., Iqbal, S. H., & Khalid, A. N. (2006). Biodiversity of Mushrooms Ectomycorrhiza. Peck. its ectomycorrhiza, a new record from Himalayan Moist Temperate Forests of Pakistan. Pakistan Journal of Botany, 38, 1271–1277.

Niazi, A. R., Khalid, A. N., & Iqbal, S. H. (2007). Descolea flavoannulata and its ectomycorrhiza from Pakistan‟s Himalayan moist temperate forests. Mycotaxon, 101, 375–383.

Niazi, A. R., Khalid, A. N., & Iqbal, S. H. (2010). New records of ectomycorrhizae from Pakistan. Pakistan Journal of Botany, 24, 4335–4343.

Nieto, M. P., & Carbone, S. S. (2009). Characterization of juvenile maritime pine ( Ait.) ectomycorrhizal fungal community using morphotyping, direct sequencing and fruitbodies sampling. Mycorrhiza, 19, 91–98.

Nilson, S., & Persson, O. (1977). Fungi of Northern Europe 2: Gill-Fungi. Penguin, p. 114.

Nuhn, M. E., Binder, M., Taylor, A. F., Halling, R. E., & Hibbett, D. S. (2013). Phylogenetic overview of the Boletineae. Fungal Biology, 117, 479–511.

O‟Donnell, K. (1979). Zygomycetes in culture, vol. 2, Palfrey contributions in botany. Department of Botany, University of Georgia, Athens, p. 257.

O'Dell, T. E., Ammirati, J. F., & Schreiner, E. G. (2000). Species richness and abundance of ectomycorrhizal basidiomycete sporocarps on a moisture gradient in the Tsuga heterophylla zone. Canadian Journal of Botany, 77, 1699–1711.

O'Dell, T. E., Massicotte, H. B., & Trappe, J. M. (1993). Root colonization of Lupinus latifolius Agardh. and Pinus contorta Dougl. by Phialocephala fortinii Wang & Wilcox. New Phytologist, 124, 93–100.

264

Osmundson, T. W., Robert, V. A., Schoch, C. L., Baker, L. J., Smith, A., Robich, G., ... & Garbelotto, M. M. (2013). Filling gaps in biodiversity knowledge for macrofungi: contributions and assessment of an herbarium collection DNA barcode sequencing project. PLoS One, 8, e62419.

Overholts, L. O. (1919). Some Colorado Fungi. Mycologia, 11, 245–258.

Palfner, G., & Agerer, R. (1998a). Balsamia alba Harkness + Pinus jeffreyi Grev. and Balf. Descriptions of Ectomycorrhizae, 3, 1–6.

Palfner, G., & Agerer, R. (1998b). Leucangium carthusianum + Pseudotsuga menziesii. Descriptions of Ectomycorrhizae, 3, 37–42.

Peay, K. G., Kennedy, P. G., & Bruns, T. D. (2008). Fungal community ecology: a hybrid beast with a molecular master. Bioscience, 58, 799–810.

Peay, K. G., Kennedy, P. G., Davies, S. J., Tan, S., & Bruns, T. D. (2010). Potential link between plant fungal distributions in a dipterocarp rainforest: community phylogenetic structure of tropical ectomycorrhizal fungi across a plant soil ecotone. New Phytologist, 185,529–542.

Pegler, D. N. (1986). Agaric flora of Sri Lanka. Her Majesty's Stationary Office, London.

Peter, M., Ayer, F., & Egli, S. (2001). Nitrogen addition in a Norway spruce stand altered macromycete sporocarp production and below‐ground ectomycorrhizal species composition. New Phytologist, 149, 311–325.

Phillips, R. (2006). Mushrooms. Pan MacMillan, pp. 107–270.

Pielou, E. C. (1969). An introduction to mathematical ecology. Wiley-Interscience, pp. 1– 286.

Pielou, E. C. (1975). Ecological diversity. Wiley Interscience, pp. 1–159.

Porras-Alfaro, A., Herrera, J., Natvig, D. O., Lipinski, K., & Sinsabaugh, R. L. (2011). Diversity and distribution of soil fungal communities in a semiarid grassland. Mycologia, 103, 10–21.

265

Porter, T. M., Shokralla, S., Baird, D., Golding, G. B., & Hajibabaei, M. (2016). Ribosomal DNA and plastid markers used to sample fungal and plant communities from wetland soils reveals complementary biotas. PloS One, 11, e0142759.

Rawat, A., Singh, A., Singh, A. B., Gaur, S. N., Kumar, L., Roy, I., & Ravidrun, P. (2000). Clinical immunologic evaluation of Cedrus deodara Pollen: a new allergen from India. Allergy, 55, 620–626.

Razaq, A. (2013). Molecular characterization and identification of gilled fungi from Himalayan moist temperate forests of Pakistan using internal transcribed spacers (ITS) of rDNA. PhD thesis, Department of Botany, University of the Punjab, Lahore.

Rinaldi, A. C., Comadini, O., & Kuyper, T. W. (2008). Ectomycorrhizal fungal diversity: separating the wheat from the chaff. Fungal Diversity, 33, 1–45.

Rudawska, M., Kieliszewska-Rokicka, B., Leski, T., & Oleksyn, J. (1995). Mycorrhizal status of a Scots pine (Pinus sylvestris L.) plantation affected by pollution from a phosphate fertilizer plant. Water, Air, and Soil Pollution, 85, 1281–1286.

Saba, M., Ahmad, I., & Khalid, A. N. (2015). New reports of Inocybe from pine forests in Pakistan. Mycotaxon, 130, 671–681.

Sanmee, R., Tulloss, R. E., Lumyong, P., Dell, B., & Lumyong, S. (2008). Studies on Amanita (Basidiomycetes: ) in Northern Thailand. Fungal Diversity, 32, 97–123.

Sarnari, M. (1998). Monographia illustrata del genre Russula in Europa. Associazione Micologica Bresadola, Trento.

Sarwar, S. Khalid, A. N., Hanif, M., & Niazi, A. R. (2012). Suillus flavidus and its ectomycorrhizae with Pinus wallichiana in Pakistan. Mycotaxon, 12, 225–232.

Sarwar, S., Hanif. M., Khalid, A. N., & Guinberteau, J. (2011). Diversity of Boletes in Pakistan – Focus on Suillus brevipes and Suillus sibiricus. Proceedings of the 7th International Conference on Mushroom Biology Mushroom Products (ICMBMP7). 123–133.

266

Sarwar, S., Jabeen, S., & Khalid, A. N. (2013). Additions to ectomycorrhizae associated with Populus ciliata Wall Ex. Royle from Pakistan. Journal of Yeast and Fungal Research, 4, 26–32.

Sarwar, S., Saba, M., Khalid, A. N., & Dentinger, B. M. (2015). Suillus marginielevatus, a new species and S. triacicularis, a new record from Western Himalaya, Pakistan. Phytotaxa, 203, 169‒177.

Schmidt, P. A., Bálint, M., Greshake, B., Bandow, C., Römbke, J., & Schmitt, I. (2013). Illumina metabarcoding of a soil fungal community. Soil Biology and Biochemistry, 65, 128–132.

Schoch, C. L., Seifert, K. A., Huhndorf, S., Robert, V., Spouge, J. L., Levesque, C. A., ... & Miller, A. N. (2012). Nuclear ribosomal internal transcribed spacer (ITS) region as a universal DNA barcode marker for Fungi. Proceedings of the National Academy of Sciences, 109, 6241–6246.

Schoch, C. L., Sung, G. H., López-Giráldez, F., Townsend, J. P., Miadlikowska, J., Hofstetter, V., ... & Gueidan, C. (2009). The Ascomycota tree of life: a phylum-wide phylogeny clarifies the origin and evolution of fundamental reproductive and ecological traits. Systematic Biology, 58, 224–239.

Schramm, J. R. (1966). Plant colonization studies on black wastes from anthracite mining in Pennsylvania. Transactions of the American Philosophical Society, 56, 5–189.

Seif, E., Leigh, J., Liu, Y., Roewer, I., Forget, L., & Lang, B. F. (2005). Comparative mitochondrial genomics in zygomycetes: bacteria-like RNase P RNAs, mobile elements and a close source of the group I intron invasion in angiosperms. Nucleic Acids Research, 33, 734–744.

Seifert, K. A. (2009). Progress towards DNA barcoding of fungi. Molecular Ecology Resources, 9, 83–89.

Seifert, K., Morgan-Jones. G., Gams, W., & Kendrick, B. (2011). The genera of Hyphomycetes. CBS-KNAW Fungal Biodiversity Centre, Utrecht, pp. 1–997.

267

Seress, D., Dima, B., & Kovács, G. M. (2015). Characterisation of seven Inocybe ectomycorrhizal morphotypes from a semiarid woody steppe. Mycorrhiza, 26, 215– 225.

Sesli, E. (2007). Preliminary checklist of macromycetes of the East and Middle Black Sea regions of Turkey. Mycotaxon, 99, 71–74.

Shaffer, R. L. (1972). North American of the Subsection Foetentinae. Mycologia, 64, 1008–1053.

Sharma, B. M., & Singh, B. M. (1990). Ectomycorrhizal fungi associated with different forest trees of Himachal Pradesh. Conference paper, Current trends in mycorrhizal research. Proceedings of the National Conference on Mycorrhiza, Haryana Agricultural University, Hisar, India, pp. 13–15.

Shi, L., Guttenberger, M., Kottke, I., & Hampp, R. (2002). The effect of drought on mycorrhizas of (Fagus sylvatica L.): changes in community structure, and the content of carbohydrates and nitrogen storage bodies of the fungi. Mycorrhiza, 12, 303–311.

Shimono, Y., Kato, M., & Takamatsu, S. (2004). Molecular phylogeny of Russulaceae (Basidiomycetes; Russulales) inferred from the nucleotide sequences of nuclear large subunit rDNA. Mycoscience, 45, 303–316.

Shinde, U.A., Phadke, A. S., Nair, A. M., Mungantiwar, A. A., Dikshit, V. J., & Saraf, M. N. (1999). Preliminary studies on the anti-inflammatory analgesic activity of Cedrus deodara (Roxb.) Loud. wood oil. Journal of Ethnopharmacology, 65, 21–27.

Siddiqui, M. F., Shaukat, S. S., Ahmed, M., Khan, N., & Khan, I. A. (2013). Vegetation- Environment relationship of conifer dominating forests of moist temperate belt of Himalayan and Hindukush regions of Pakistan. Pakistan Journal of Botany, 45, 577– 92.

Simpson, D. P. (1979). Cassell's Latin Dictionary (5th edn.). London, Cassell Ltd., p. 883.

268

Singer, R. (1986). The Agaricales in modern taxonomy (4th edn). Koeltz Scientific Books, Koenigstein, pp. 1–981.

Singh, L., & Lakhanpal, T. N. (2000). Growth ectomycorrhiza development in Cedrus deodara seedlings inoculated with different vegetative inoculum formulations. Journal of Mycology and Plant Pathology, 30, 64–67.

Šmilauer, P., & Lepš, J. (2014). Multivariate Analysis of Ecological Data using CANOCO 5 (2nd ed.). Cambridge University Press.

Smith, M. E., Douhan, G. W., & Rizzo, D. M. (2007). Ectomycorrhizal community structure in a xeric Quercus wood based on rDNA sequence analysis of sporocarps pooled roots. New Phytologist, 174, 847–863.

Smith, J. E., Molina, R., Huso, M. M., Luoma, D. L., McKay, D., Castellano, M. A., ... & Valachovic, Y. (2002). Species richness, abundance, and composition of hypogeous and epigeous ectomycorrhizal fungal sporocarps in young, rotation-age, and old- growth stands of Douglas-fir (Pseudotsuga menziesii) in the Cascade Range of Oregon, USA. Canadian Journal of Botany, 80, 186–204.

Smith, M. E., Henkel, T. W., Aime, M. C., Fremier, A. K., & Vilgalys, R. (2011). Ectomycorrhizal fungal diversity and community structure on three co-occurring leguminous canopy tree species in Neotropical rain forest. New Phytologist, 192, 699–712.

Smith, M. E., Henkel, T. W., Uehling, J. K., Fremier, A. K., Clarke, H. D., & Vilgalys, R. (2013). The ectomycorrhizal fungal community in a Neotropical forest dominated by the endemic dipterocarp Pakaraimaea dipterocarpacea. PloS One, 8, e55160.

Smith, S. E., & Read, D. J. (2008). Mycorrhizal Symbiosis, (3rd edn.). Academic Press Elsevier, London.

Stangl, J. (1989). Die gattung Inocybe in Bayern. Hoppea, 46, 5–388.

269

Stoyke, G., & Currah, R. S. (1993). Resynthesis in pure culture of a common subalpine fungus-root association using Phialocephala fortinii and Menziesia ferruginea (). Arctic and Alpine Research, 25, 189–193.

Stucki, B., & Khan, H. A. (1999). Working plan for Utror-Desan forests of Kalam Forest Division. Peshawar, Pakistan Nizam Printing Press.

Stuntz, D. E. (1947). Studies in the genus Inocybe. I. New and noteworthy species from Washington. Mycologia, 39, 21–55.

Sugiyama, J. (1998). Relatedness, phylogeny, and evolution of the fungi. Mycoscience, 39, 487–51.

Sundberg, W., & Bessette, A. (1987). Mushrooms: A Quick Reference Guide to Mushrooms of North America (Macmillan Field Guides). New York, Collier Books, p. 20.

Talbot, P. H. B. (1971). Principles of Fungal Taxonomy. The Macmillan Press, London, pp. 1–274.

Tamura, K., Stecher, G., Peterson, D., Filipski, A., & Kumar, S. (2013). MEGA6: molecular evolutionary genetics analysis version 6.0. Molecular Biology and Evolution, 30, 2725–2729.

Tanabe, Y., O'Donnell, K., Saikawa, M., & Sugiyama, J. (2000). Molecular phylogeny of parasitic Zygomycota (Dimargaritales, Zoopagales) based on nuclear small subunit ribosomal DNA sequences. Molecular Phylogenetics and Evolution, 16, 253–262.

Tanabe, Y., Saikawa, M., Watanabe, M. M., & Sugiyama, J. (2004). Molecular phylogeny of Zygomycota based on EF-1α and RPB1 sequences: limitations and utility of alternative markers to rDNA. Molecular Phylogenetics and Evolution, 30, 438–449.

Tanabe, Y., Watanabe, M. M., & Sugiyama, J. (2002). Are Microsporidia really related to Fungi?: a reappraisal based on additional gene sequences from basal fungi. Mycological Research, 106, 1380–1391.

270

Tanabe, Y., Watanabe, M. M., & Sugiyama, J. (2005). Evolutionary relationships among basal fungi (Chytridiomycota and Zygomycota): Insights from molecular phylogenetics. The Journal of General and Applied Microbiology, 51, 267–276.

Tandan, S. K, Chandra, S., Gupta, S., & Lal, J. (1998). Pharmacodynamic effects of Cedrus deodara wood essential oil. Indian Journal of Pharmaceutical Sciences, 60, 20–23.

Taylor, A. F. S. (2002). Fungal diversity in ectomycorrhizal communities: sampling efforts and species detection. Plant and soil, 198, 77–84.

Taylor, D. L., & Bruns, T. D. (1999). Community structure of ectomycorrhizal fungi in a Pinus muricata forest: minimal overlap between the mature forest and resistant propagule communities. Molecular Ecology, 8, 1837–1850.

Taylor, D., Booth, M. G., McFarland, J. W., Herriott, I. C., Lennon, N. J., Nusbaum, C., & Marr, T. G. (2008). Increasing ecological inference from high throughput sequencing of fungi in the environment through a tagging approach. Molecular Ecology Resources, 8, 742–752.

Taylor, J. W., Jacobson, D. J., Kroken, S., Kasuga, T., Geiser, D. M., Hibbett, D. S., & Fisher M. C. (2000). Phylogenetic species recognition and species concepts in fungi. Fungal Genetics and Biology, 31, 21–32.

Tedersoo, L., Abarenkov, K., Nilsson, R. H., Schüssler, A. Grelet, G. A., Kohout, P.,... & Kõljalg, U. (2011). Tidying up international nucleotide sequence databases: ecological, geographical sequence quality annotation of ITS sequences of mycorrhizal fungi. PloS One, 6, e24940.

Tedersoo, L., Hansen, K., Perry, B. A., & Kjøller, R. (2006). Molecular morphological diversity of pezizalean ectomycorrhiza. New Phytologist, 170, 581–596.

Tedersoo, L., Kõljalg, U., Hallenberg, N., & Larsson, K. H. (2003). Fine scale distribution of ectomycorrhizal fungi and roots across substrate layers including coarse woody debris in a mixed forest. New Phytologist, 159, 153–165.

271

Tedersoo, L., May, T. W., & Smith, M. E. (2010). Ectomycorrhizal lifestyle in fungi: global diversity, distribution, evolution of phylogenetic lineages. Mycorrhiza, 20, 217–263.

Tedersoo, L., Naadel, T., Bahram, M., Pritsch, K., Buegger, F., Leal, M., ... & Põldmaa, K. (2012). Enzymatic activities and stable isotope patterns of ectomycorrhizal fungi in relation to phylogeny and exploration types in an afrotropical rain forest. New Phytologist, 195, 832–843.

Tedersoo, L., Suvi, T., Jairus, T., Ostonen, I., & Põlme, S. (2009). Revisiting ectomycorrhizal fungi of the genus Alnus: differential host specificity, diversity and determinants of the fungal community. New Phytologist, 182, 727–735.

Thiers, H. D. (1975). California Mushrooms: A Field Guide to the Boletes. New York, NY, Hafner Press, p. 45.

Trappe, J. M. (1962). Fungus associates of ectotrophic mycorrhizae. Botanical Review, 28, 538–606.

Trevor, E. Y., Egger, K. N., & Peterson, L. R. (2001). Ectendomycorrhizal associations– characteristics and functions. Mycorrhiza, 11, 167–177.

Tulloss, R. E. (2005). Amanita velosa (Peck) Lloyd. Tulloss Amanita website: http://www.amanitaceae.org/

Tulloss, R.E., & Lindgren, J.E. (1994). Amanita novinupta - a rubescent, white species from the Western United States and Southwestern Canada. Mycotaxon, 51, 179–190.

UNEP. (1998). Land cover assessment and monitoring Pakistan (Vol. 10A). Environment Assessment Programme for Asia and the Pacific, Bangkok. pp. 1–50.

Vaario, L. M., Xing, S. T., Xie, Z. Q., Lun, Z. M., Sun, X., & Li, Y. H. (2006). In situ in vitro colonization of Cathaya argyrophylla (Pinaceae) by ectomycorrhizal fungi. Mycorrhiza, 16, 137–142.

Väre, H., Vestberg, M., & Eurola, S. (1992). Mycorrhiza and root-associated fungi in Spitsbergen. Mycorrhiza, 1, 93–104.

272

Verma, B., & Reddy, M. S. (2015). Suillus indicus sp. nov. (Boletales, Basidiomycota), a new boletoid fungus from northwestern Himalayas, India. Mycology, 6, 35–41.

Vesterholt, J. (1989). A revision of Hebeloma sect. Indusiata in the Nordic countries. Nordic Journal of Botany, 9, 289–319.

Visser, S. (1995). Ectomycorrhizal fungal succession in jack pine stands following wildfire. New phytologist, 129, 389–401.

Vizzini, A. (2015). Nomenclatural novelties. Index Fungorum, 244.

Waite, S. (2000). Statistical ecology in practice. A guide to analysing environmental and ecological field data. Harlow, England, Prentice Hall.

Walker, J. F., Miller Jr, O. K., & Horton, J. L. (2008). Seasonal dynamics of ectomycorrhizal fungus assemblages on oak seedlings in the southeastern Appalachian Mountains. Mycorrhiza, 18, 123–132.

Wang, B., & Qiu, Y. L. (2006). Phylogenetic distribution evolution of mycorrhizas in plants. Mycorrhiza, 16, 299–363.

Wang, C. J. K., & Wilcox, H. E. (1985). New species of ectendomycorrhizal and pseudomycorrhizal fungi: Phialophora finlandia, Chloridium paucisporum, and Phialocephala fortinii. Mycologia, 77, 951–958.

Wang, Q., & Guo, L. D. (2010). Ectomycorrhizal community composition of Pinus tabulaeformis assessed by ITS-RFLP and ITS sequences. Botany, 88, 590–595.

Wang, Q., Gao, C., & Guo, L. D. (2011). Ectomycorrhizae associated with Castanopsis fargesii (Fagaceae) in a subtropical forest, China. Mycological Progress, 10, 323– 332.

Watling, R., & Lee, S. S. (1995). Ectomycorrhizal fungi associated with members of the Dipterocarpaceae in peninsular Malaysia. Journal of Tropical Forest Science, 7, 657– 669.

273

Weiss, M., Selosse, M. A., Rexer, K. H., Urban, A., & Oberwinkler, F. (2004). : a hitherto overlooked cosm of heterbasidiomycetes with a broad mycorrhizal potential. Mycological Research, 108, 1003–1010.

White, M. M. (2002). Taxonomic and molecular systematic studies of the Harpellales (Trichomycetes) toward understanding the diversity, evolution and dispersal of gut fungi. Doctoral dissertation, University of Kansas Press, p. 172.

White, M. M. (2006). Evolutionary implications of a rRNA based phylogeny of Harpellales. Mycological Research, 110, 1011–1024

White, M. M., Cafaro, M. J., Gottlieb, A. M. (2001). Taxonomy and systematics of Trichomycetes-past, present and future. In: Misra, J. K., & Horn, B. W. (eds.) Trichomycetes and other fungal groups. Science Publishers, Enfield, pp. 27–37.

White, T. J., Bruns, T. D., & Taylor, L. J. (1990). Amplification direct sequencing of fungal ribosomal RNA genes for Phylogenetics. In: Innis, M. A., Gelf, D. H., Sninsky, J. J., & White, T. J. (eds.). PCR protocols: A guide to methods applications. Academic Press, New York, pp. 315–322.

Whittaker, R. H. (1972). Evolution and measurement of species diversity. Taxon, 1, 213–251.

Wilcox, H. E., & Wang, C. J. K. (1987). Mycorrhizal and pathological associations of dematiaceous fungi in roots of 7-month-old tree seedlings. Canadian Journal of Forest Research, 17, 884–899.

Wolff, R. L., Lavialle, O., Pedrono, F., Pasquier, E., Delue, L. G., Marpean, A. M., & Aitzelmuller, K. (2001). Fatty acid composition of Pinaceae as taxonomic markers. Lipids, 36, 439–451.

Wurzburger, N., Bidartondo, M. I., & Bledsoe, C. S. (2001). Characterization of Pinus ectomycorrhizas from mixed conifer and pygmy forests using morphotyping and molecular methods. Canadian Journal of Botany, 79, 1211–1216.

Yamada, A., & Katsuya, K. (1996). Morphological classification of ectomycorrhizas of Pinus densifiora, Mycoscience, 37, 145–155.

274

Yousaf, N., Kreisel, H., & Khalid, A. N. (2013). Bovista himalaica sp. nov. (gasteroid fungi; Basidiomycetes) from Pakistan. Mycological Progress, 12, 569–574.

Zak, J. C. (1973). Classification of Ectomycorrhizae. In: Marks, G. C., & Kozlowaski, T. T. (eds.), Ectomycorrhizae, their ecology physiology. Academic Press, New York, pp. 43–73.

Zambonelli, A., Iotti, M. Amicucci, A., & Pisi, A. (1999). Anatomical and morphological characterization of mycorrhizae of Tuber maculatum Vittad. and Ostrya carpinifolia Scop. Micologia Italiana, 3, 29–35.

Zambonelli, A., Pisi, A., & Tibiletti, A. (1997). Anatomical and morphological characterization of mycorrhizae of Tuber indicum Cooke Masee with Pinus pinea L. and Quercus cerris L. Micologia Italiana, 1, 29–36.

Zeitlmayr, L. (1976). Wild Mushrooms: An Illustrated Handbook. Garden City Press, Hertfordshire, pp. 93–94.

Zhang, J., Takahashi, K., Kono, Y., Suzuki, Y., Takeuchi, S., Shimizu, J., Yamaguchi, I., Chijimatsu, M., & Sakurai, A. (1990). Bioactive condensed tannins from bark: Chemical properties enzyme inhibition anti plant viral activities. Nippon Noyaku Gakkaishi, 15, 585–591.

Zhang, L., Yang, J., & Yang, Z. (2004). Molecular phylogeny of eastern Asian species of Amanita (Agaricales, Basidiomycota): taxonomic and biogeographic implications. Fungal Diversity, 17, 219–238.

Zhang, T., Wang, N. F., Liu, H. Y., Zhang, Y. Q., & Yu, L. Y. (2016). Soil pH is a key determinant of soil fungal community composition in the Ny-Ålesund region, Svalbard (High Arctic). Frontiers in Microbiology, 7, 227.