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The effect of grazing on arbuscular mycorrhizal fungi in temperate grasslands van der Heyde, Mieke Elisabeth 2016

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THE	EFFECT	OF	GRAZING	ON	ARBUSCULAR		MYCORRHIZAL	FUNGI	IN	TEMPERATE	GRASSLANDS	by	Mieke	Elisabeth	van	der	Heyde	B.Sc.,	McGill	University,	2013	A THESIS SUBMITTED IN PARTIAL  FULFILLMENT OF THE REQUIREMENTS FOR THE DEGREE OF  Master of Science  in THE COLLEGE OF GRADUATE STUDIES Biology UNIVERSITY OF BRITISH COLUMBIA (Okanagan) September 2016 © Mieke Elisabeth van der Heyde, 2016	The College of Graduate Studies The undersigned certify that they have read, and recommend to the College of Graduate Studies for acceptance, a thesis entitled: submitted by  in partial fulfilment of the requirements of the degree of  . Supervisor, Professor (please print name and faculty/school above the line)Supervisory Committee Member, Professor (please print name and faculty/school in the line above)Supervisory Committee Member, Professor (please print name and faculty/school in the line above) University Examiner, Professor (please print name and faculty/school in the line above) External Examiner, Professor (please print name and university in the line above)   (Date Submitted to Grad Studies) Additional Committee Members include: (please print name and faculty/school in the line above)(please print name and faculty/school in the line above)Page 1 of 1 Revised: October-15-14 ii	ABSTRACT	Managed	grazing,	involving	large	animals	destined	for	human	consumption,	covers	more	than	25%	of	the	land	surface	and	has	the	capacity	to	alter	ecosystems,	often	leading	to	desertification,	woody	encroachment,	and	deforestation	(Asner	et	al	2014).	Arbuscular	Mycorrhizal	(AM)	fungi		are	ubiquitous	root	symbionts	that	colonize	80%	of	terrestrial	plants	and	influence	plant	productivity	and	community	composition.	Despite	the	importance	of	AM	fungi	for	plant	communities,	the	effect	of	grazing	on	AM	fungal	communities	is	largely	unknown.		I	used	grazing	exclosures	of	varying	ages	to	compare	AM	fungal	community	and	infectivity	in	grazed	and	ungrazed	plots,	as	well	as	several	environmental	variables	that	may	be	affected	by	grazing.	AM	fungal	community	composition	was	not	significantly	different	between	grazed	and	ungrazed,	but	grazing	increased	spore	density	while	decreasing	soil	hyphal	length.	This	may	be	attributed	to	the	plasticity	of	AM	fungi	in	response	to	environmental	conditions,	flexibility	allowed	isolates	to	respond	to	grazing	without	shifting	community	composition.	None	of	the	environmental	variables	was	related	to	the	change	in	AM	fungi,	indicating	that	variables	not	measured	may	be	responsible.	Time	since	the	exclosure	was	established	was	the	only	variable	related	to	community	dissimilarity	between	grazed	and	ungrazed.	As	the	age	of	the	exclosure	increased,	the	dissimilarity	also	increased,	highlighting	the	importance	of	long-term	studies	in	furthering	our	understanding	of	grazer-fungi	interactions.			iii	PREFACE	This	study	was	conducted	using	grazing	exclosures	established	and	maintained	by	the	Ministry	of	Forests,	Lands,	and	Natural	Resource	Operations	in	the	Southern	Interior	British	Columbia,	Canada.	Sampling	was	conducted	with	permission	and	assistance	from	Range	Ecologist	Dr.	Rick	Tucker,	who	provided	the	plant	community	data.	I	designed	the	experiment	and	performed	the	fieldwork	with	the	help	of	members	of	the	Hart	lab.	I	was	responsible	for	processing	the	samples	and	performing	molecular	work.	The	BC	Ministry	of	Environment,	Technical	Services	Lab	(Victoria,	British	Columbia)	did	the	soil	chemical	analysis,	and	IBEST	Genomics	Resources	Core	at	the	University	of	Idaho	did	the	sequencing.	I	performed	the	sequence	analysis	and	collaborated	with	Dr.	Jonathan	Bennet	on	the	data	analysis.	I	wrote	my	thesis	with	the	guidance	of	my	supervisor	Dr.	Miranda	Hart,	and	it	was	reviewed	by	my	supervisory	committee:	Dr.	David	Scott	and	Dr.	Jason	Pither	from	the	University	of	British	Columbia	(Okanagan).	iv	TABLE	OF	CONTENTS	ABSTRACT…………………………………………………………………………….. ii PREFACE………...……………………………………………………………………. iii TABLE OF CONTENTS……...………………………………………………………. iv LIST OF TABLES……………………..……………………………………………… vii LIST OF FIGURES…………………….…………………………………………….. viii LIST OF ABBREVIATIONS………………………………………………………….. x ACKNOWLEDGMENTS……………………...……………………………………… xi CHAPTER 1 : INTRODUCTION...……...……………………………………………. 1	1.1. Grasslands	grazing	and	Arbuscular	Mycorrhizal	(AM)	fungi…..……….. 1 1.2. Ambiguous	effects	of	grazing	on	AM	fungi...………………………………. 2	1.3. Response	to	herbivory	depends	on	response	variable………………….. 3	1.3.1. AM	fungal	infectivity..…...…………………………………………….. 4 1.3.2. AM	fungal	community	structure.……………………………………. 5	1.4. Herbivory	response	depends	on	ecological	context………………………7	1.4.1. AM	identity	affects	responses	to	herbivory…………………………7	1.4.2. Host	plant/	plant	community	influence	the	effect	of	herbivory.…8	1.4.3. Soil	Chemistry	………….……………………………………………….9	1.4.4. Time	………………………………………………………………….……9	2.1. Study	Site	..……………………………………………………………….…...13 2.2. Sampling…………………………………………………………………….…131.5. Research	Objectives.…………………………………………………………10	CHAPTER 2: METHODS…………...………...…………………………………………13 v	2.2.1. Soil…….…………………………………………….……………….…..13	2.2.2. AM	fungal	infectivity….………………………………………….…….14	2.2.3. Soil	Chemistry……………………………………………………….…15	2.2.4. Plant	Community....……………………………………………………15	2.3. Community	Analysis………………………………………………………….16	2.3.1. Molecular	Analysis…………………………………………………….16	2.3.2. Sequence	Analysis…..…………………………………………………17	2.4. 	Data	Analysis….………………………………………………………….......18	2.4.1. Does	grazing	affect	the	AM	fungal	community? .………………….18	2.4.1.1. AM	fungal	infectivity	(%	root	col,	spore	density,	hyphal	length)	responses	to	grazing..………………………………18	2.4.1.2. AM	fungal	community	responses	to	grazing..………………18	2.4.2. Does	grazing	affect	the	mycorrhizal	environment?	……………...19	2.4.3. Are	changes	in	AM	fungal	community	related	to	grazing	induced	changes	in	soil	environment	and/or	age	of	exclosure? ……….……20	CHAPTER	3:	RESULTS………………………………………………………........……..21        3.1. Does	grazing	affect	the	AM	fungal	community? ………………………….213.1.1. AM	fungal	infectivity	responses	to	grazing..……………………….21	3.1.2. AM	fungal	community	responses	to	grazing...……….……………21	3.2. Does	grazing	affect	the	mycorrhizal	environment?	……….….…………21	3.2.1. Plant	and	soil	responses	to	grazing………….………….…………..22	3.3. Are	changes	in	AM	fungal	community	related	to	grazing	induced	changes	in	soil	environment	and/or	age	of	exclosure? ….……………..24 vi	CHAPTER 4: DISCUSSION...………………….……………………….....……...……26     4.1. Does	grazing	affect	AM	fungal	infectivity?……………...……..……………26 4.2. Does	 grazing	 affect	 AM	 fungal	 community	 composition?…………..…….29 4.3. Does	grazing	affect	the	mycorrhizal	environment?		Plantand	soil	responses	to	grazing……..………...…………………..…………...31	4.4. The effect of time…………………………………….…..……………………..35 CHAPTER 5: CONCLUSION...….………………………………........……………….37   5.1. AM fungal infectivity…...………………………...………………………….….37 5.2. AM fungal community..…...………………………………………………….…37 5.3. The effect of time………...…………………………………………………………38BIBLIOGRAPHY…….......……………………………………………….……………..46APPENDIX A: SUPPLEMENTARY MATERIAL...………………………………….58vii	LIST	OF	TABLES	Table	1:	Description	of	the	nine	exclosures	selected.	………………………………………... 42Table	2:	Biotic	and	abiotic	responses	to	grazing	at	nine	grazing	exclosures.	Differences	in	plant	and	soil	variables	between	grazed	and	ungrazed	were	tested	using	Wilcoxon	signed	rank	tests.	95%	confidence	intervals	than	did	not	overlap	with	0	and	p-values	less	than	0.1	are	bolded	as	an	indication	that	there	was	some	difference	between	grazed	and	ungrazed.	…………………………………………………………... 42Table	3:	The	change	in	percent	cover	with	grazing	of	the	most	abundant	plant	species.	Only	plants	that	were	found	in	most	sites	(at	least	7/9)	and	made	up	a	significant	proportion	(at	least	10%)	of	the	plant	community	were	included.	Change	was	calculated	as	=	(%	cover	in	grazed)-(%cover	in	ungrazed)	…………………………... 43Table	4:	Blocked	indicator	species	analysis	of	AMF	Families	and	isolates	as	indicators	of	grazed	or	ungrazed	plots.	Only	Families	with	more	than	2	isolates	were	included	in	the	family	analysis.	Only	the	nine	strongest	indicator	isolates	were	shown.	………………………………………………………………………………………………………………44 Table	5:	Summary	of	Multimodel	Inference	Statistics.	Italicized	descriptions	are	the	dependent	values	for	the	global	models.	Model	summaries	are	in	the	Appendix	......45viii	LIST	OF	FIGURES	Figure	1:	The	standardized	change	in	AMF	infectivity	with	grazing.	Change	=	(infectivity	in	grazed)	–	(infectivity	in	ungrazed).	The	solid	red	line	indicates	no	difference	between	grazed	and	ungrazed.	Open	circles	show	the	change	at	each	site,	closed	circles	represent	the	standardized	median,	and	error	bars	show	the	95%	confidence	intervals.	Col	=	Colonization,	Hyph	=	Hyphal	length,	Spore	=	Spore	density,	Sur	=	surface	soil,	Sub	=	subsurface	soil.		P-values	of	the	paired	Wilcox	signed	rank	test	were	as	follows:	Col	Sur	(p-value=0.930),	Hyph	Sub	(p-value	=	0.160),	Hyph	Sur	(p-value	=	0.039),	Spore	Sub	(p-value=0.359),	Spore	Sur(p-value=0.055).	N=9		…………………………………………………………………………………………….. 38Figure	2:	Nonmetric	multidimensional	scaling	(NMDS)	showing	distance	between	AMF	communities	from	grazed	and	ungrazed	plots	for	both	soil	depths	(data	pooled).	Ordination	axes	were	calculated	using	Bray-Curtis	distance	metric	(Bray	and	Curtis	1957),	with	4	axes	and	a	stress	value	of	0.1746.		Grazing	was	not	a	significant	factor	any	of	the	axes.	An	ordination	of	each	layer	separately	also	finds	no	significant	effect	of	grazing	on	any	of	the	axes.	……………………………………………………39Figure	3:	The	standardized	change	in	AMF	richness	and	evenness	with	grazing.	The	solid	red	line	indicates	no	difference	between	grazed	and	ungrazed.	Open	circles	show	the	change	at	each	site,	closed	circles	represent	the	standardized	mean,	and	error	bars	show	the	95%	confidence	intervals.	Sur	=	surface	soil,	Sub	=	subsurface	ix	soil.	P-values	for	the	paired	Wilcox	signed	rank	test	were	as	follows:	Evenness	Sub	(p-value=0.301),	Evenness	Sur	(p-value	=	0.735),	Richness	Sub	(p-value	=	0.875),	Richness	Sur	(p-value	=	0.938).	N=9	……………………………………………………………....... 40Figure	4:	The	relationship	between	Age	of	the	exclosures	and	the	dissimilarity	between	AMF	communities	in	grazed	and	ungrazed.	Plot	A	represents	the	surface	communities	and	plot	B	shows	the	subsurface	communities.	Surface	r=0.66,	subsurface	r=0.64.	……………………………………………………………………………...…………... 411	CHAPTER	1:	GENERAL	INTRODUCTION	1.1 Grasslands	grazing	and	Arbuscular	Mycorrhizal	(AM)	fungi	Grassland	plants	have	a	long	evolutionary	history	with	herbivores	that	allows	them	to	tolerate	high	levels	of	grazing	(Stebbins	1981).	 Humans	depend	on	this	coevolution	to	maintain	pastures	that	are	essential	for	food	production	across	many	cultures.	Managed	grazing,	which	involves	large	animals	destined	for	human	consumption,	covers	more	than	25%	of	the	global	land	surface	and	can	result	in	desertification,	woody	encroachment,	and/or	deforestation	(Asner	et	al.	2004).	 As	an	economically	and	ecologically	important	human	impact,	it	is	vital	we	understand	all	the	factors	influencing	the	productivity	and	biodiversity	of	these	systems,	including	Arbuscular	Mycorrhizal	(AM)	fungi.	AM	fungi	are	soil	microbes	that	colonize	the	roots	of	plants	and	form	a	symbiotic	partnership	with	their	hosts.	These	organisms	make	up	the	phylum	Glomeromycota	and	colonize	80%	of	terrestrial	plants	(Smith	and	Read	2008).	They	are	beneficial	fungi	capable	of	acquiring	nutrients	for	the	plant	(Smith	and	Read	2008),	protecting	them	from	pathogens	(Sikes	et	al.	2009),	and	helping	plants	deal	with	stress	such	as	drought	(reviewed	in	Augé	2001)	and	salt	stress	(Sharifi	et	al.	2007).	 In	return,	plants	supply	mycorrhizal	fungi	with	carbon,	without	which	the	fungi	cannot	survive	(Pearson	and	Jakobsen	1993).	If	the	plant	is	carbon	limited	(i.e.	from	losing	productive	biomass	to	herbivores)	then	there	is	also	less	carbon	available	for	the	fungi.	Such	grazing-induced	limits	on	the	AM	symbiosis	may	have	far	reaching	effects.	AM	fungi	are	important	in	ecosystems,	capable	of	influencing	plant	biodiversity	and	productivity	(van	der	Heijden,	Bardgett,	and	van	Straalen	2008;	Köhl,	Oehl,	and	van	2	der	Heijden	2014).		This	is	particularly	true	in	grasslands	because	virtually	all	perennial	plants	in	these	systems	form	AM	symbioses.	Despite	the	importance	of	AM	fungi	in	grasslands,	little	is	known	about	the	effect	of	grazing	on	AM	fungi	in	these	systems.	While	the	plant	response	to	grazing	is	relatively	well	understood,	the	AM	fungal	response	is	not.	 Since	grazing	imposes	changes	in	carbon-	supply	by	removing	productive	biomass,	it	may	be	a	selective	force	on	AM	fungal	communities	as	well.	1.2 Ambiguous	effects	of	grazing	on	AM	fungi	There	has	been	considerable	research	on	AM	fungi	and	grazing,	but	AM	fungal	responses	to	grazing	are	inconsistent	and	even	contradictory	among	studies	(Hokka	et	al.	2004;	Klironomos	et	al.	2004;	Mikola	et	al.	2005).	This	lack	of	consensus	is	not	due	lack	of	research:		there	have	been	58	papers	in	this	area	since	1980,	not	including	reviews	and	meta-analyses.	 Rather,	the	problem	lies	in	the	controversy	over	the	effect	of	herbivory	on	the	AM	symbiosis	One	theory	has	been	proposed	to	explain	the	response	of	AM	fungi	to	herbivory	primarily	in	terms	of	carbon	supply,	the	carbon	limitation	hypothesis	(Wallace	1987;	Gehring	and	Whitham	1994).	The	carbon	limitation	hypothesis	predicts	that	herbivory	should	decrease	fungal	colonization	overall	as	host	limits	the	amount	of	carbon	available	to	root	symbionts	(Wallace	1987).	Overall,	there	is	mixed	support	for	the	carbon	limitation	hypothesis	in	the	literature,	with	considerable	support	among	different	systems	and	hosts	(Gehring	and	Whitham	1994;	Frank	et	al.	2003;	Wardle	et	al.	2002;	Barber	et	al.	2012)	Many	studies,	however,	show	herbivory	elicits	either	no	response	in	root	colonization		 3		(Pietikäinen	et	al.	2005),	or	increased	root	colonization	 (Eom	et	al.	2001;	Hokka	et	al.	2004;	Wearn	and	Gange	2007;	Keitaro	et	al.	2012;	Techau	et	al.	2004;	Nishida	et	al.	2009)	or	variable	colonization	responses	(Bethlenfalvay	and	Dakessian	1984;	Hokka	et	al.	2004;	Klironomos	et	al.	2004;	Piippo	et	al.	2011).	A	recent	meta-analysis	challenging	the	carbon	limitation	hypothesis	(Barto	and	Rillig	2010)	found	little	evidence	for	reduction	in	root	colonization	following	herbivory,	and	suggested	that	other	factors	may	determine	fungal	response	to	grazing.	Considering	that	grazing	presents	more	that	simply	a	carbon	limitation	to	AM	fungi	may	help	to	better	understand	disturbance	effects	on	AM	fungi	and	resulting	plant	communities.		In	addition	to	carbon	limitation,	grazing	imparts	a	suite	of	changes,	which	can	affect	plant,	fungus,	and	soil	that	may	impinge	upon	the	AM	symbiosis.	 These	include	soil	compaction	(Wallace	1987),	increases	in	Nitrogen	(N)	availability	from	root	exudates	(Hamilton	et	al.	2008),	increased	nutrient	availability	through	feces	and	urine	of	herbivores	(Huntly	1991)	or	plant	mediated	N-mineralization	(Holland	and	Detling	1990;	Hamilton	et	al.	2008),	and/or	changes	to	plant	communities	(Wikeem	et	al.	2012).	 Given	that	each	of	these	factors	may	affect	AM	partners	differentially,	understanding	the	response	of	the	AM	symbiosis	to	grazing	may	be	difficult	using	only	single	response	variables.				1.3 Response	to	herbivory	depends	on	response	variable		 AM	fungi	may	be	affected	by	grazing	in	multiple	ways,	including	(but	not	limited	to)	 changes	 in	 infectivity	 of	 root	 and	 soil,	 and	 changes	 in	 fungal	 community	composition.	 Most	 studies,	 however,	 tend	 to	 gauge	 fungal	 response	 using	 only	 one	variable,	typically	percent		 4	root	length	colonization.	By	neglecting	to	measure	multiple	response	variables,	important	grazing	responses	may	be	overlooked.	1.3.1 AM	fungal	infectivity		 Percent	root	colonization,	a	measure	of	relative	AM	fungi	abundance,	is	the	most	commonly	reported	response	of	AM	fungi	to	grazing,	and	also	perhaps	the	most	variable.	Although	reduced	percent	root	colonization	following	herbivory	is	predicted	by	the	carbon	limitation	hypothesis,	and	is	commonly	reported	(Gehring	and	Whitham	1994;	Barber	et	al.	2012;	Saravesi	et	al.	2014)	there	are	many	examples	where	herbivory	leads	to	increased	root	colonization	(Eom	et	al.	2001;	Kula	et	al.	2005;	Nishida	et	al.	2009;	Wearn	and	Gange	2007;	Keitaro	et	al.	2012).	Increases	in	AM	fungal	colonization	following	defoliation	may	be	a	response	to		 increased	resources	associated	with	grazing	(i.e.	through	urine	and	feces	of	grazing	herbivores	(Baron	et	al.	2001)),	as	plants	may	opportunistically	invest	more	heavily	in	AM	fungi	to	exploit	additional	nutrients	and	stimulate	regrowth	(Barto	and	Rillig	2010).	Counterintuitively,	increases	in	AM	colonization	could	result	from	increased	carbon	availability	as	defoliated	plants	briefly	increase	carbon	exudation	in	the	roots	(Holland	et		al.	1996;	Bazot	et	al.	2005;	Hamilton	et	al.	2008).	 This	pulse	of	carbon	may	be	reflected	in	increases	in	fungal	abundance	overall,	including	hyphal	length,	root	colonization,	and	spore	density.	Such	‘pulses’	of	resources	belowground	may	explain	why	the	increases	in	colonization	have	been	shown	to	decrease	after	two	weeks	(Nishida	et	al.	2009).	It	may	also	explain	why	only	low	frequency	herbivory	is	associated	with	increases	in	root	colonization:	if	the	frequency	of	defoliation	is	too	high,	the	plant	has	no	carbon	to	“pulse”	in	the	roots	and	the	colonization	decreases	(Klironomos	et	al.		 5	2004).	In	addition	to	biological	explanations,	changes	in	AM	fungal	colonization	after	grazing	may	be	related	to	the	measure	itself.	 Percent	root	length	colonized	may	over-	or	underestimate	fungal	investment	into	roots	and	does	not	account	for	differences	among	fungal	structures	such	as	storage	vesicles,	intra-radical	hyphae,	and	arbuscules	(Hart	and	Reader	2002).	Soil	fungal	structures	such	as	extra-radical	hyphae	and	spores	are	also	ignored.	Thus,	AM	fungal	investment	may	be	reduced	following	defoliation,	but	not	perceived	by	%	root	length	colonized.	Similarly,	as	root	biomass	is	almost	uniformly	reduced	by	herbivory	(Holland	et	al.	1996;	Thornton	et	al.	2016;	Ferraro	and	Oesterheld	2002),	total	fungal	colonization	may	decrease,	even	if	there	are	no	changes,	or	increases	in	%	root	colonization.	 The	more	grazing	reduces	the	root	length,	the	more	the	absolute		 amount	of	fungal	colonization	will	be	overestimated.		1.3.2 AM	fungal	community	structure		 Herbivory	may	also	result	in	changes	to	AM	fungal	communities.	Since	herbivory	changes	the	plant	community	through	grazing	(Frank	2005),	soil	compaction	(Shelton	et	al.	2014),	and	changes	to	soil	fertility	(Frank	and	Groffman	1998),	these	changes	may	translate	into	changes	below	ground,	as	AM	fungi	and	plant	communities	are	tightly	linked	(van	der	Heijden	et	al.	1998).	As	with	other	response	variables,	there	is	no	clear	pattern	to	grazing	response.	AM		 fungal	diversity	has	been	shown	to	both	decline	(Eom	et	al.	2001;	Su	and	Guo	2007;	Ba	et	al.	2012),	and	increase	in	response	to	grazing	(Ba	et	al.	2012).	While	fewer	in	number,	there	are	also	studies	showing	compositional	changes	(Eom	et	al.	2001;		 6	Murray	et	al.	2010;	Yang	et	al.	2013),	but	not	in	all	(Shelton	et	al.	2014).	One	explanation	for	varying	results	is	the	degrees	of	grazing	intensity;	Ba	et	al	(2012)	found	increases	in	diversity	at	light	grazing,	but	reductions	at	higher	grazing	levels.	 Thus,	compositional	changes	may	only	be	apparent	at	high	herbivory.	Compositional	changes	as	a	result	of	grazing	suggest	that	there	are	different	levels	of	grazing	tolerance	between	AM	fungi.	Where	AM	fungal	diversity	was	reduced	with	grazing,	the	AM	fungi	under	grazing	were	predominantly	Glomeraceae	isolates	(Eom	et	al.	2001;	Su	and	Guo	2007;	Ba	et	al.	2012).	 The	fungi	in	this	family	have	fast	growth	rates	(de	la	Providenicia	et	al.	2005;	Hart	and	Reader	2005),	abundant,	early	sporulation	(Oehl	et	al.	2009;	Pringle	and	Bever	2002)	and	are	often	considered	disturbance	tolerant.	This	suggests	the	Glomeraceae	may	be	the	most	grazing	tolerant,	but	compositional	changes	because	of	grazing	do	not	always	result	in	increases	in	Glomeraceae	(Murray	et	al.	2010;	Yang	et	al.	2013).	Glomeraceae	can	be	sensitive	to	grazing	(Su	and	Guo	2007;	Ba	et	al.	2012)	and	non-Glomeraceae	can	increase	with	grazing	(Eom	et	al.	2001;	Murray	et	al.	2010).	The	variability	in	response	may	be	an	artifact	of	community	measurements;	most		published	reports	used	spore	sievings	to	quantify	AM	fungal	communities	(Eom	et	al.	2001;	Su	and	Guo	2007;	Murray	et	al.	2010;	Ba	et	al.	2012;	Shelton	et	al.	2014).	Studies	using	spores	to	identify	communities	are	limited	to	the	soil,	and	spores	producing	AM	fungi	only,	possibly	missing	changes	in	AM	fungi	community.	Only	two	studies	have	used	a	molecular	approach	(Yang	et	al.	2013;	Guo	et	al.	2016).	 Of	those,	Yang	et	al	(2013)	found	that	grazing	changed	the	communities	in	the	roots	only	and	Guo	et	al		 7	(2016)	found	that	responses	depended	on	the	steppe	type	and	plant	communities.	It	is	well	established	that	different	sample	types	(ie	root,	soil,	spores,	sequences,	etc.)	result	in	significantly	different	AM	fungal	communities	(Hempel	et	al.	2009;	Selosse	et	al.	2016).	Thus	both	techniques	and	the	nature	of	the	sample	may	contribute	to	variation	in	observed	responses.	1.4 Herbivory	response	depends	on	ecological	context		 Even	if	studies	were	able	to	measure	the	full	suite	of	AM	fungal	responses,	it	is	likely	that	the	discord	among	studies	would	exist,	albeit	reduced.	 The	context	of	grazing		(i.e.	plant	identity,	AM	fungal	identity,	soil	chemistry)	may	be	as	important	as	response	variable	in	determining	disturbance	outcomes	and	may	significantly	alter	the	ability	to	detect,	or	change	the	direction	of	grazing	response.	1.4.1 AM	identity	affects	responses	to	herbivory		 If	AM	taxa	have	differential	tolerance	to	conditions	associated	with	grazing,	then	it	is	likely	that	AM	fungal	community	composition	will	affect	grazing	outcomes.	 While	there	are	many	reports	in	the	literature	about	differential	disturbance	tolerance	among	the	Glomeromycota	(Cuenca	et	al.	1998;	Jansa	et	al.	2002;	Hart	and	Reader	2004;	Schnoor	et	al.	2011),	we	have	yet	to	fully	understand	how	this	variation	is	organized	(i.e.	whether	or	not		it	is	phylogenetically	conserved	(Hart	and	Reader	2004)	or	a	result	of	selection	pressure	(Maherali	and	Klironomos	2012)).	In	terms	of	grazing,	such	variation	could	result	in	positive	(if	fungi	are	disturbance		 tolerant)	or	negative	feedback	(if	fungi	are	disturbance	intolerant)	between	host	and	fungi.	There	is	only	indirect	evidence	of	this	feedback	in	the	literature:	we	know	that	AM	fungi	differ	in	their	capacity	to	improve	compensatory	growth	capacity	(Bennett	et		 8	al.	2009;	Bennett	and	Bever	2007),	or	ability	to	stimulate	herbivore	defense	(Nishida	et	al.	2009;	Barber	2013),	but	there	is	yet	no	direct	evidence	for	differential	grazing	tolerance	among	AM	hosts	attributable	to	AM	identity.	Due	to	the	high	levels	of	functional	variation	among	AM	fungal	isolates	(Hart	and	Reader	2002b;	Munkvold	et	al.	2004;	Koch	et	al.	2004)	it	is	not	yet	possible	to	determine	which	fungi,	and	under	which	contexts,	AM	fungi	would	lead	to	positive	versus	negative	feedback	for	defoliated	hosts.	1.4.2 Host	plant/	plant	community	influence	the	effect	of	herbivory		 Varying	reports	of	AM	fungal	grazing	response	may	also	stem	from	differences	in	host	identity.	 There	is	a	history	of	research	showing	that	response	to	herbivory	is	a	continuum	among	plant	taxa,	with	some	intolerant,	and	others	that	are	competitively	dominant	under	grazing	(Ellison	1960;	Robertson	1971;	Crawley	1990).	If	hosts	are	differentially	tolerant	of	grazing,	then	it	follows	that	fungi	associated	with	tolerant	hosts	should	be	less	affected	by	grazing.	For	example,	the	root	biomass	of	grasses	is	more	resistant	to	change	following	defoliation	than	annual	plants	(reviewed	in	Ferraro	and	Oesterheld	2002),	which	may	explain	why	percent	root	colonization	for	annual	plants	decreases	more	than	grasses	and	forbs	(Barto	and	Rillig	2010).	If,	disturbance	tolerance	among	plants	results	from	compensatory	growth	(McNaughton	1983),	this	may	also	stimulate	AM	fungal	activity	in	order	to	acquire	the	necessary	nutrients.	However,	if	plant	grazing	tolerance	is	due	to	plant	traits	conferring	stress	tolerance	(eg.	quiescence	during	grazing,	perenniating	structures	below	ground	(Huhta	et	al.	2009),	or	unpalatable	tissues	(Diaz	et	al.	2007),	AM	fungal	activity	may	be	reduced	(quiescence),	or	be	unchanged	(unpalatable	plants)	in	response	to	grazing.				 9		1.4.3 Soil	Chemistry		 Soil	chemistry	can	mask	the	effects	of	herbivory	on	AM	fungal	colonization,	(Techau	et	al.	2004;	Ruotsalainen	and	Eskelinen	2011)	plant	benefit	(Aguilar-Chama	and	Guevara	2012)	and	mycorrhizal	community	(Murray	et	al.	2010).	 This	is	not	surprising	as	not	only	are	plants	better	able	to	tolerate	herbivory	when	there	are	abundant	resources	(McNaughton	1983),	but	high	soil	fertility	is	also	known	to	suppress	AM	fungi	(Treseder	and	Allen	2002).		Simply	put,	there	are	situations,	particularly	at	high	nutrient	availability,	where	AM	fungi	respond	to	soil	fertility	more	than	of	herbivory	(Techau	et	al.	2004),	and	this	may	be	compounded	by	AM	fungal	specific	differences	in	their	tolerance	to	soil	nutrients	(Murray	et	al.	2010).	 In	such	cases,	herbivory	effects	may	only	be	apparent,	or	may	change	directions,	under	different	nutrient	regimes.	Other	soil	variables	like	pH	can	reverse	the	effect	of	herbivory	entirely		 (Ruotsalainen	and	Eskelinen	2011)	possibly	by	affecting	the	plant	and	AM	fungal	community	composition.	Routslainen	and	Eskilinen	(2011)	showed	grazing	reduced	root	colonization	only	in	acidic	soil,	while	in	ungrazed	soil,	root	colonization	increased	with	acidity.	 Thus,	studies	showing	changes	in	AM	fungal	community	composition	(Murray	et	al.	2010;	Ba	et	al.	2012)	may	be	the	result	of	low	pH	rather	than	grazing	itself.	Understanding	how	soil	chemistry	interacts	with	herbivory	is	vital	to	determining	whether	the	AM	fungal	responses	general	responses	to	herbivory	or	due	to	concomitant	changes	in	soil	conditions.	1.4.4 Time		 Time	is	also	very	important	and	little	understood	factor	in	determining	AM	fungal	responses	to	herbivory.	Some	AM	fungal	responses,	such	as	percent	colonization,		 10	can	occur	almost	instantly	but	may	be	equally	short	lived	(Nishida	et	al.	2009).	 Others,	such	as	changes	in	community	composition	may	take	years	to	develop	and	years	to	recover	from	(Shelton	et	al.	2014).		Variation	among	published	studies	in	time	frame,	both	between	the	herbivory	event	and	the	AM	fungal	measurement,	but	also	in	the	length	of	time	the	plants	were	exposed	to	herbivory	treatments,	may	significantly	change	the	outcome.	Perhaps	it	takes	time	to	affect	the	AM	fungal	community	because	grazing	also	takes	time	to	affect	the	soil	condition	and	plant	community,	and	possibly	decades	for	them	to	recover	(Pyke	et	al.	2016;	Krzic	et	al.	2012;	Shelton	et	al.	2014).				1.5 Research	Objectives		 My	goal	is	to	determine	how	grazing	affects	the	AM	fungal	community	in	temperate	grasslands.	The	literature	on	grazing	and	AM	fungi	is	inconsistent	due	to	several	factors	such	as	host	identity,	AM	identity,	sampling	bias,	and	ecological	context.	To	address	these	shortcomings	I	measured	multiple	indicators	of	AM	fungal	infectivity,	including	community	composition.	While	I	could	not	completely	control	for	ecological	context,	I	was	able	to	measure	several	soil	and	plant	variables	to	understand	what	changes	in	response	to	grazing	and	whether	these	changes	affect	the	AM	fungal	communities.	I	used	an	established,	long	term	grazing	exclosure	chronosequence,	to	answer	the	following	questions:				 1) Does	grazing	affect	AM	fungal	infectivity	(root	colonization,	spore	density,	and	hyphal	length),	and/or	AM	fungal	community	composition?	• I	predicted	that	AM	fungal	infectivity	would	remain	the	same	as	grasses		 11	and	forbs	are	habituated	to	grazing	and	are	able	to	maintain	their	symbionts.	• However,	I	expected	there	would	be	shifts	in	AM	fungal	community	composition	as	the	grazed	and	ungrazed	plots	are	under	different	selection	pressures.	2) Does	grazing	affect	the	mycorrhizal	environment?	That	is,	does	grazing	affect	factors	known	to	influence	the	AM	fungal	community	such	as	soil	chemistry,	soil	density,	plant	community,	and	plant	biomass?	I	predicted	that	grazing	would	reduce	plant	biomass	and	shift	the	plant	community	by	selecting	for	more	grazing	tolerant	species.	o I	predicted	that	grazing	would	alter	the	chemical	profile	of	the	soil	because	grazing	may	also	increase	nutrient	availability	through	defecation/urination.	o Finally,	I	predicted	grazing	would	lead	to	increased	soil	compaction	compared	to	soil	in	exclosures	due	to	the	activity	of	herbivores.				3) Are	changes	in	AM	fungal	infectivity/community	related	to	grazing	induced	changes	in	soil	environment	and/or	age	of	the	exclosure?	• I	predicted	changes	in	AM	fungal	infectivity	would	be	related	to	changes	in	soil	nutrient	availability	and/or	plant	biomass,	as	those	variables	affect	how	much	carbohydrates	are	available	for	the	AM	fungi.	• I	predicted	changes	in	plant	community	and	soil	nutrients	to	be	the	strongest	predictors	for	AM	fungal	community	composition	because	both	are	known	to	cause	shifts	in	community	composition	through	host	preference	and	nutrient	stress.		 12	• I	also	predicted	the	differences	between	grazed	and	ungrazed	would	increase	with	the	age	of	the	exclosure,	as	the	communities	would	have	more	time	to	differentiate.		 13	CHAPTER	2:	METHODS					2.1 Study	Site		 I	selected	nine	grazing	exclosures	established	by	the	Ministry	of	Forests,	Lands,	and	Natural	Resource	Operations	in	the	Southern	Interior	British	Columbia,	Canada.	Sites	were	selected	based	on	similar	vegetation	types	(grasslands),	elevation	(~1100m),	and	grazing	(cattle	and	horses).	Seven	of	the	sites	were	located	in	the	Hamilton	Mountain	area	about	25-30km	east	of	Merritt,	BC.	The	remaining	two	were	located	near	Tunkwa	Provincial	Park,	BC.	Soils	at	all	sites	were	loam	Black	Chernozems	that	developed	from	glacial	till	deposits		of	volcanic	rock	and	limestone	(Young	et	al	1992).	The	area	has	a	semi-arid	climate	with	hot,	dry	summers.	Each	exclosure	was	fenced	off	using	barbed	wire	fencing	to	prevent	any	grazing	within.	The	exclosures	varied	in	age	from	17	to	85	years	(See	Table	1).	Each	exclosure	ranged	from	0.5-1	ha,	and	were	paired	with	a	grazed	site	directly	adjacent.				2.2 Sampling		2.2.1 Soil		 Samples	were	collected	in	May	2015,	every	10	m	along	a	70m	transect	(10m	from	the	fence).	Twenty	soil	samples	at	each	site	(10	grazed,	10	exclosure)	were	taken,	5	at	the	surface	(0-15cm)	and	5	at	the	subsurface	(15-30cm).	 In	total,	I	collected	9	sites	x	2	grazing	treatments	x	2	depths	x	5	=	180	samples	to	be	used	for	AM	fungal	analyses.	To	collect	subsurface	samples,	I	first	dug	a	small	hole	to	15	cm	and	then	used	a	soil	corer	to	collect	deeper	soil.	The	samples	were	kept	cool	in	the	field	until	they	could	be	frozen,	at	most	48	hours	from	time	of	collection.	They	were	dried	for	two	days	at	60	°C,	homogenized	with	500µm	sieve,	and	subsampled	for	DNA	extraction.	Then	samples	were		 14	pooled	by	plot	because	there	was	not	enough	soil	to	quantify	the	soil	chemistry	and	mycorrhizal	infectivity	of	each	sample	separately.	By	pooling	the	samples	I	was	able	to	get	values	for	the	plot	as	a	whole,	but	not	the	heterogeneity	within	plot.	In	all,	36	samples	from	18	grazed/ungrazed	plots	were	used	to	determine	AM	fungal	abundance	and	soil	properties.	In	June	2015	further	sampling	was	undertaken	to	determine	soil	bulk	density.	 Five		undisturbed	cores	were	taken	in	each	plot	near	the	original	samples,	except	they	were	5m	from	the	fence	instead	of	10m.	The	samples	were	taken	closer	to	the	fence	in	June	to	minimise	the	destructive	effect	of	biomass	harvesting	and	core	sampling	on	the	integrity	of	the	exclosures,	and	their	future	uses.	Again,	cores	were	taken	at	the	surface	(0-15cm)	and	subsurface	 (15-30cm).	The	samples	were	dried	for	five	days	at	40	°C	and	weighed.		As	one	site	(Muscrat)	was	too	rocky	for	coring,	I	ended	up	with	160	bulk	density	cores	that	were	kept	separate	until	they	were	weighed.	The	average	bulk	density	of	the	five	samples	in	each	plot	was	used	as	the	plot	bulk	density	in	future	analysis.	2.2.2 AM	fungal	infectivity		 Root	segments	in	dried	soil	samples	were	randomly	selected	from	dried	soil		samples	for	staining.	This	selection	likely	covered	only	the	most	common	plant	species.	Washed	root	segments	were	cut	into	1cm	pieces	and	stored	in	50%	ethanol	before	being	stained	with	Chlorazol	Black	E	(Brundrett	et	al	1984).	Percent	colonization	was	determined	using	the	magnified	intersection	method	(McGonigle	et	al	1990)	to	determine	what	proportions	of	the	roots	were	colonized	by	AM	fungi.	 Only	the	surface	samples	contained	enough	root	segments	to	accurately	quantify	percent	colonization.	Spores	were	extracted	from	soil	samples	using	a	wet-sieving	technique	(Klironomos	et	al.	1993)	to	calculate	spore	abundance.	Hyphal	length	was	determined	using	200	mL	of	soil	suspension	was		 15	combined	with	1	mL	of	a	mixture	of	europium	(III)	thenoyl-trifluoroacetonate	and	a	fluorescent	brightener	(Anderson	and	Westmoreland	1971).	The	suspensions	were	then	stained	for	1	hr,	and	then	filtered	through	nitrocellulose	filter	papers	using	a	50%	ethanol	wash.	The	filters	were	mounted	on	microscope	slides	and	computer-imaging	software	was	used	to	estimate	hyphal	length.	2.2.3 Soil	Chemistry		 The	pooled	soil	samples	from	each	plot	were	sent	to	the	BC	Ministry	of	Environment,	Technical	Services	Lab	(Victoria,	British	Columbia)	for	chemical	analysis.	Soil	Phosphorous	was	extracted	using	the	Bray-P1	method	 (Bray	et	al.	1945)	and	quantified	using	a	visible	spectrophotometer.	A	1:1	mixture	of	soil:water	was	used	to	measure	soil	pH	using	an	ion	meter.	Total	C	an	N	were	quantified	using	combustion	elemental	analysis.	2.2.4 Plant	Community		 The	aboveground	biomass	for	ten	0.5	m2		quadrats	per	site	was	removed,	dried	at	60°C	for	three	days	and	weighed.	In	order	to	obtain	a	better	measure	of	the	productive	biomass	in	each	plot,	only	live	biomass	was	included	and	litter	was	removed	prior	to	harvest.	Five	plots	were	harvested	inside	the	exclosures	and	the	other	five	outside,	located	adjacent	to	where	soil	samples	were	collected	and	5	m	from	the	fence	to	minimize	the	destructive	effect	on	the	study	sites.	Plant	communities	inside	and	outside	the	exclosures	were	determined	using	a		 random	number	generator	to	select	50	Daubenmire	(50	x	20cm)	quadrats	along	five	parallel	30m	transects	spaced	5m	apart.	Percent	cover	was	estimated	at	each	quadrat	for	each	plant	species	before	all	fifty	quadrats	were	combined	to	give	a	description	of	the	plant	community	of	the	plot	as	a	whole	(Daubenmire	1959).	The	Ministry	of	Forests,	Lands,	and		 16	Natural	Resource	Operations	completed	this	in	2014	for	5	sites	(Tunkwa	New,	Tunkwa	Old,	Repeater,	Muscrat,	Goose),	and	in	2015	for	4	sites	(Stipa	Rich,	Stipa	Nel,	Bluegrass,	and	Aspen).	I	assisted	with	surveys	at	Stipa	Rich,	but	most	work	was	done	by	range	ecologists	at	the	Ministry.	These	plant	surveys	are	usually	conducted	every	two	decades	to	monitor	changes	in	community,	as	differences	can	take	decades	to	develop.	Therefore	a	one-year	difference	between	sampling	should	not	be	enough	to	noticably	alter	plant	communities.				2.3 Community	Analysis		2.3.1 Molecular	analysis		 DNA	was	extracted	from	125mg	soil	following	the	protocol	of	the	FastDNA-96TM	Soil	Microbe	DNA	(MP	Biomedicals)	after	first	drying,	sieving	and	homogenizing	soil.	Nested	PCR	was	performed	on	the	samples,	first	to	amplify,	then	to	attach	barcodes	for	Illumina	sequencing.	In	the	first	PCR,	Glomeromycota	specific	primers	WANDA	(Dumbrell	et	al	2010)	and	AML2	(Lee	et	al	2007)	were	used	to	amplify	a	500bp	fragment	of	the	small	subunit	of	rDNA.	Samples	were	amplified	in	a	20	μL	reaction	consisting	of	11.75	μL	ddH20,	5	μL	5x	PCR	buffer	(Promega),	1	μL	MgCl2	(BioLabs),	0.5	μL	dNPTs	(Amresco),	0.25	μL	BSA	(BioLabs),	0.5	μL	GoTaq	(Promega),	and	0.5	μL	of	each	primer.	This	mixture	was	heated	for	2	minutes	at	95°C	then	underwent	34	cycles	of	95	°C	for	1	min,	55	°C	for	1	min,	72	°C	for	1	min,	finishing	with	72°C	for	5	minutes.	Barcodes	were	added	to	PCR	products	in	a	20	μL	reaction	of	9.2	μL	ddH20,	4	μL	5x	PCR	buffer	(Promega),	3.6	μL	MgCl2(BioLabs),	0.4	μL	dNPTs(Amresco),	0.6	μL	BSA(BioLabs),	0.2	μL	GoTaq(Promega),	and	0.5	μL	of	each	barcode.	Cycling	conditions	were:	1	min	at	95°C,	10	cycles	of	30	secs	at	95°C,	30	secs	at	60°C,	1	min	at	68°C,	and	ending	with	5	min	at	68°C.	 Samples	were	frozen	and	sent	to	the	IBEST	Genomics	Resources	Core	at	the	University	of	Idaho	for	sequencing	using	Illumina		 17	Sequencing	Technology	(Bennet	2004).	2.3.2 Sequence	Analysis		 Raw	sequences	were	processed	using	QIIME	(Caporoso	et	al.	2011).	 The	raw	sequence	files	contained	approximately	4.2	million	sequences,	3.2	million	of	which	were	retained	after	low	quality	(Q	,	3)	,	and	reads	with	more	than	3	N	characters	were	removed.	We	used	USEARCH	(Edgar	2010)	and	a	97%	identity	threshold	to	pick	Operational	Taxonomic	Units	(OTUs)	using	maarjAM	database	(Opik	et	al.	2010)	as	a	reference	database.	This	resulted	in	28927	OTUs,	which	was	reduced	to	351	OTUs	after	filtering	out	those	comprising	less	than	0.01%	of	the	data	set.	Representative	sequences	of	these	OTUs	were	aligned	using	MUSCLE	(Edgar	2004),	then	BLASTed	(Alschul	et	al.	1990)	against	both	the	maarjAM	database	(Opik	et	al.	2010)	and	the	NCBI	database	(Geer	et	al.	2010)	to	ensure	consistent	taxonomic	assignment.	In	total,	11	non-Glomeromycotan	OTUs	were	removed,	and	OTUs	matching	identical	MaarjAM	virtual	taxon	accessions	were	merged	to	give	a	final	number	of	62	OTUs	with	an	average	of	14000	sequences	per	sample.	The	resulting	OTU	table	was	rarefied	to	1718	sequences/sample,	as	that	was	the	lowest	sequencing	depth	for	all	samples.	The	final	table	was	log	transformed	before	analysis	because	of	the	skewed	distribution	of	sequences.		 		 18			2.4 Data	Analysis	Data	analysis	was	done	using	R	v3.2.1	(R	Core	Team	2015).	Because	of	my	limited	sample	size,	I	am	considering	p-values	of	less	than	0.1	significant.	2.4.1 Does	grazing	affect	the	AM	fungal	community?		2.4.1.1	 AM	fungal	infectivity	(%	root	col,	spore	density,	hyphal	length)	responses	to	grazing	I	tested	the	effect	of	grazing	on	each	AM	fungal	infectivity	variable	using	the	paired	Wilcox	signed	rank	test	for	non-parametric	data	from	the	R	package	MASS	(Venables	and	Ripley	2002).	The	samples	were	paired	by	site	comparing	infectivity	values	at	grazed	and	ungrazed.	The	soil	layers	were	tested	separately,	as	they	were	not	independent,	and	not	paired	for	this	test.									2.4.1.2	 AM	fungal	community	responses	to	grazing		 To	determine	if	there	were	differences	in	AM	fungal	community	composition	resulting	from	grazing,	I	used	non-metric	multidimensional	scaling	in	the	R	package	‘vegan’	to	create	an	ordination	based	on	the	Bray	Curtis	distance	metric	(Oksanen	et	al	2015).	The	dimensionality	of	the	ordination	was	increased	until	model	stress	dropped	below	0.2	–	the	resultant	model	had	4	axes	with	a	stress	of	(0.1745).	We	tested	each	axis	separately	for	response	to	grazing	using	a	variation	on	the	split	plot	mixed	model	above.	However,	as	we	took	multiple	samples	per	plot,	we	also	included	a	random	factor,	representing	sample	as	a	repeated	measure	within	plot.	Note	that	because	these	tests	are	not	independent,	alpha	should	be	adjusted	from	0.1	to	0.1/3=0.033	for	statistical	significance.	For	subsequent	AM	fungal	community	analyses,	we	pooled	the	data	by	plot.	We	then	calculated	AM	fungal	richness	and	evenness	(Pielou	1966)	and	tested	the	effect	of	grazing		 19	using	the	paired	Wilcox	signed	rank	test.	In	addition,	we	conducted	indicator	specie	analysis	(Dufresne	and	Legendre	1997)	to	identify	AM	fungal	families	and	isolates	that	were	indicators	of	grazing	using	‘indicspecies’	in	R	(De	Caceres	and	Legendre	2009).	2.4.2 Does	grazing	affect	the	mycorrhizal	environment?	Plant	and	soil	responses	to	grazing	The	soil	and	plant	variables	measured	were	tested	with	the	same	paired	Wilcox	signed	rank	tests	with	the	two	soil	layers	being	tested	separately.		Are	changes	in	AM	fungal	community	related	to	grazing	induced	changes	in	soil	environment	and/or	age	of	exclosure?	To	determine	how	environmental	changes	and	age	of	exclosure	might	affect	the	AM		 fungal	community	response	to	grazing,	I	compared	environmental	variables	to	the	changes	in	AM	fungal	infectivity	and	community.	I	calculated	the	change	in	AM	fungal	infectivity,	soil	chemistry,	and	plant	community	as:	Change	in	infectivity	=infectivity	in	grazed	plots-infectivity	in	ungrazed.		 I	used	a	multi-model	inference	approach	in	the	R	package	MuMin	(Barton	2016)	to	tests	all	possible	combinations	of	the	predictors	(changes	in	environmental	variables	and	age	of	exclosure)	on	changes	in	spore	density,	and	hyphal	length,	and	then	ranked	the	resultant	models	by	AICc	scores,	which	represent	model	fit	penalized	by	the	number	of	parameters	(Burnham	and	Anderson	2002).	Variable	weights	were	subsequently	derived	from	the	model	weights	as	the	sum	of	AICc	scores	for	all	models	in	which	that	variable	appeared.	Only	variables	with	the	highest	weights	were	plotted.	Multicollinearity	of	predictors	were	tested	using	the	variance	inflation	factor	in	the	R	package	‘car’	(Fox	and	Weisberg	2011).		 20	To	test	whether	changes	in	environmental	variables	or	exclosure	age	explain	changes	in	AM	fungal	community	composition,	we	calculated	the	Bray-Curtis	dissimilarity	between	Hellinger	transformed	(Legendre	and	Gallagher	2001)	grazed	and	ungrazed	plots	at	each	site.	Then	we	used	the	same	multi-model	inference	approach	introduced	previously	to	determine	which	grazing	induced	changes	in	the	soil	and	plant	are	responsible	for	the	changes	AM	fungal	community	composition,	and	how	age	of	exclosure	ranks	as	a	predictor.	21		CHAPTER	3:	RESULTS					3.1 Does	grazing	affect	the	AM	fungal	community?		3.1.1 AM	fungal	infectivity	responses	to	grazing		Percent	Root	Colonization		 Grazing	did	not	affect	percent	root	colonization	(p=0.930)	95%	CI	[-16.0,	10.5],	(Figure	1).	 Average	percent	colonization	across	all	plots	was	31.8	(SE	3.12)	Spore	density		 Spore	density	tended	to	be	higher	under	grazing	than	in	the	exclosure	at	the	surface	(p	=	0.054)	95%	CI	[-8.0,	236.5]	(Figure	1).	The	subsurface	soil	showed	less	of	a	response	(p	=	0.359)	95%	CI	[-71.0,	169].	At	the	surface,	ungrazed	soil	had	an	average	spore	density	of	194.9	spores/10g	(SE	41.71),	while	the	average	in	grazed	soil	was	302.6	spores/10g	(SE	26.15).	In	the	B	layer,	spore	density	was	lower,	with	an	average	of	108	spores/10g	(SE	32.19)	in	ungrazed	and	156	spores/10g	(SE	26.90)	under	grazing.	Soil	hyphal	length		 Grazing	is	associated	with	reduced	soil	hyphal	length	in	the	surface	soil	(p=0.039)	95%	CI	[-1.85,	-0.05]	(Figure	1).	In	the	subsurface,	there	was	no	association	with	grazing	(p=0.160)	95%	CI	[-1.9,	0.3].	Average	hyphal	length	in	the	surface	soil	was	3.26	mm/g	(SE	0.27)	in	ungrazed	and	2.32	mm/g	(SE	0.24)	under	grazing,	while	in	the	subsurface	average	hyphal	length	was	1.84	mm/g	(SE	0.31)	in	ungrazed	and	0.99	mm/g	(SE	0.133).	3.1.2 AM	fungal	Community	responses	to	grazing		AM	fungi	Alpha	diversity	Neither	AM	fungal	richness	(surface	p=0.938,	subsurface	p=0.875)	nor	AM	fungal	evenness	(surface	p=0.734,	subsurface	p=0.301)	appears	affected	by	grazing	(Figure	2).	AM	22		fungal	richness	was	35.8	(SE	1.07)	at	the	surface	and	30.1	(SE	1.23).	Average	evenness	was	0.678	(SE	0.012)	at	the	surface	and	0.643	(SE	0.018)	in	the	subsurface	soil.	AM	fungal	community	changes		 Across	all	sites,	we	identified	62	OTUs	from	8	families:	2	Ambisporaceae,	4		 Archeaosporaceae,	3	Acaulosporaceae,	7	Diversisporaceae,	1	Pacisporaceae,	8		 Claroideoglomeraceae,	33	Glomeraceae,	3	Paragomeraceae,	and	1	Glomeromycotan	OTU	that	did	not	match	any	AM	fungal	family	in	the	databases.	There	was	no	significant	effect	of	grazing	on	AM	fungal	community	composition	(P=	0.28,	0.45,	0.46,	0.20	for	each	axis)	(Figure	3).	The	low	marginal	R-squared	values	for	each	axis	(0.0056,	0.0048,	0.0031,	and	0.014	respectively)	demonstrate	how	little	of	the	variation	could	be	explained	by	grazing	alone.	The	conditional	R-squared	values	were	considerably	higher	(0.1208,	0.1884,	0.3111,	and	0.1696	respectively)	indicating	the	importance	of	accounting	for	site	differences.	 Several	isolates	were	significant	as	indicators	of	either	grazed	or	ungrazed	plots	(Table	3):	 two	Glomus	(p=0.0031	and	p=0.0856)	a	Paraglomus	(p=0.0817),	and	an	Ambispora	(p=0.084),	isolate	were	positively	associated	with	grazing,	while	three	Diversispora	isolates	were	found	more	often	in	ungrazed	plots	(p=0.0388,	p=0.0974,	and	p=0.1024).	None	of	the	AM	fungal	families	were	indicators	of	grazing	exposure	(Table	3).				3.2 Does	grazing	affect	the	mycorrhizal	environment?		3.2.1 Plant	and	soil	responses	to	grazing		Plant	community		 Biomass	and	percent	cover		 Aboveground	biomass	was	higher	inside	exclosures	(p	<0.001)	95%	CI	[25.52,	23		54.18].	Both	treatments	were	dominated	by	grasses,	which	had	an	average	percent	cover	of	71.6	(SE	4.9),	while	forbs	had	an	average	percent	cover	of	26.9	(SE	4.1).	Dominant	species		 (>50%	percent	cover)	were	Festuca	campestris	Rydb.	and	Poa	pratensis	L.	Other	plants	that	were	found	in	most	sites	(at	least	7	out	of	9)	include	the	grasses	Koeleria	macrantha	Ledeb.,	Stipa	nelsonii	Scribn.,	and	forbs	Achilllea	millefolium	L.,	Agoseris	glauca	Pursh.,	Aster	campestris	Nuttall.,	Astragalus	miser	Hook.,	Geum	triflorum	Pursh,	Taraxacum	officinale	L.,	and	Tragopogon	dubius	Scop.	Alpha	diversity		 Plant	richness	was	unchanged	by	grazing	(21.4	plants	(SE	0.71)	in	grazed	and	18.5	plants	(SE	1.89)	in	ungrazed	plots	(P=0.44).	 Plant	evenness,	however,	was	significantly	higher	in	grazed	plots	0.592	(SE	0.053)	compared	to	ungazed	plots	0.458	(SE	0.050)	(p=0.048)	(Table	2).	Differences	in	evenness	were	largely	due	the	reduced	the	abundance	of	Festuca	campestris		in	grazed	plots	(by	89.3	%	(Goose)	50%	(Repeater,	Muscrat,	Tunkwa	Old)	or	25%	(Aspen,	Stipa	Nel,	Tunkwa	New)(See	Table	3).	One	site	(Bluegrass)	had	no	Festuca	campestris	inside	or	outside	the	exclosure.	Festuca	campestris	was	identified	as	an	indicator	of	ungrazed	plots	(P=0.0081)	in	our	indicator	species	analysis,	and	Tragopogon	dubius	was	an	indicator	of	grazing	(P=0.011).	Soil	chemistry		 Phosphorous	tended	to	be	a	little	higher	under	grazing	for	both	surface	(Adjusted	p=0.37)	and	subsurface	(Adjusted	p=0.27)	soil	layers	(See	Table	2	for	details).	Total	nitrogen	was	also	higher	under	grazing	at	the	surface	(p=0.195)	but	not	in	the	subsurface	(p=1.000).	Although	p-values	do	not	indicate	significance	because	they	had	to	be	adjusted	for	multiple	comparison,	there	seems	to	be	a	trend	towards	higher	soil	nitrogen	and	phosphorous	under	grazing.	Total	Carbon	was	not	significantly	different	under	grazing	at	24		the	surface	(p=0.31)or	subsurface	(p=1.00)	neither	was	pH	(surface	p=1.00,	subsurface	p=1.00).		Soil	density	was	also	higher	under	grazing	in	the	surface	soil	(p=0.039),	with	an	average	of	0.818	g/cc	(SE	0.0188)	under	grazing	and	0.676	g/cc	(SE	0.0314)	in	the	exclosures.	In	the	subsurface,	there	was	no	significant	difference	between	grazed	an	ungrazed	(p=1.00).				3.3	Are	changes	in	AM	fungal	community	related	to	grazing	induced	changes	in	soil	environment	and/or	age	of	exclosure?	Because	percent	root	colonization,	subsurface	hyphal	length,	and	subsurface	spore		 density	did	not	change	with	grazing;	they	were	not	included	in	this	analysis.		Spore	Density		 The	multi-model	inference	approach	to	identify	which	environmental	variable	was	related	to	the	change	in	spore	density	identified	plant	evenness	as	the	variable	with	the	most	weight.	As	plant	evenness	increased,	so	did	spore	density	(standardized	estimate	=	0.313,	weight=0.53).	However,	as	the	standard	error	(0.381)	is	greater	than	the	estimate,	and	the	95%	CI	[-0.196,	1.375]	overlaps	with	0,	this	relationship	is	not	significant	(Table	5).	Hyphal	length		 Similar	to	spore	density,	none	of	the	changes	in	environmental	variables	was	significantly	related	to	the	changes	in	hyphal	length	with	grazing.	Age	of	exclosure	was	the	most	important	variable	explaining	changes	in	soil	hyphal	length	(estimate	=	0.118,	weight=0.31).	Again,	the	standard	error	(0.265)	was	greater	than	the	estimate	and	the	95%	CI	[-0.468,	1.220]	overlaps	with	0,	indicating	little	importance	of	this	variable	AM	fungal	community	composition		 Time	since	exclosure	was	the	most	important	variable	explaining	differences	25		between	the	AM	fungal	communities	inside	and	outside	the	exclosures.	This	is	true	for	both	the	surface	(estimate=0.526,	weight=0.73)	and	the	subsurface	(estimate=0.604,	weight=0.81).	 With	increasing	age	of	the	exclosure,	the	dissimilarity	between	grazed/ungrazed	AM	fungal	communities	increased	(Fig	4).	The	correlation	coefficient	was	0.66	for	the	surface	and	0.64	in	the	subsurface.	No	other	variables	contributed	to	variation		 among	AM	fungal	communities.	26		CHAPTER	4:	DISCUSSION					 In	this	study	I	showed	that	grazing	affects	AM	fungal	infectivity	more	than	community	composition.	Further,	a	single	measure	of	AM	fungal	infectivity	is	not	sufficient	to	determine	AM	fungal	responses	to	grazing,	as	different	measures	gave	contrasting	results.	Despite	significant	changes	in	soil	fertility,	plant	community,	and	soil	density,	none	of	these	factors	were	related	to	changes	in	AM	fungal	infectivity	or	AM	fungal	community.	Rather,	the	most	the	most	important	driver	of	changes	in	AM	fungal	communities	was	time,	as	dissimilarity	between	grazed	and	ungrazed	plots	increased	over	time.				4.1 Does	grazing	affect	AM	fungal	infectivity?		Percent	Colonization		 As	predicted,	grazing	did	not	affect	percent	root	colonization.	This	result	is	consistent	with	the	literature	for	field	experiments,	which	find	either	no	change	(Bethlenfalvay	and	Dakessian	1984;	Pietikäinen	et	al.	2005;	Yang	et	al.	2013)	or	an	increase	in	percent	colonization	(Eom	et	al.	2001;	Kula	et	al.	2005).	Studies	which	show	decreases	in	percent	colonization	with	herbivory	(Barber	et	al.	2012;	Saravesi	et	al.	2014)	typically	occur	only	under	greenhouse	conditions.	 While	it	has	been	thought	that	carbon	limitation	from	the	plant	following	herbivory	would	result	in	a	decrease	in	fungal	abundance		(Gehring	and	Whitham	1994),	this	is	not	always	the	case,	especially	for	fungi	associated	with	grasslands,	which	are	adapted	to	grazing	(Barto	and	Rillig	2010).	In	these	systems,	plants	may	be	able	to	maintain	the	same	level	of	colonization,	through	defoliation	induced	carbon	exudates	(Hamilton	et	al.	2008),	compensatory	photosynthesis	in	residual	foliage	27			(Piippo	et	al.	2011)	or	through	the	action	of	a	secondary	metabolite	which	could	stimulate	mycorrhization	(Landgraf	et	al.	2012).	Alternatively,	while	I	was	unable	to	detect	a	change	in	fungal	colonization	among	grazing	treatments,	this	may	be	due	to	the	methods	used	to	estimate	fungal	abundance.	Percent	colonization	is	a	relative	measure,	thus	if	root	biomass	is	reduced	by	grazing	and	percent	colonization	remains	the	same,	there	would	be	a	net	reduction	in	total	fungal	colonization.		While	I	was	not	able	to	measure	root	biomass	in	this	study,	grazing	is	known	to	reduce	root	biomass	(Ferraro	and	Oesterheld	2002),	and	I	observed	much	less	dense	root	mats	under	grazing.		Therefore,	it	is	possible	that	while	plants	were	able	to	maintain	relative	colonization	levels	in	roots,	the	total	abundance	of	fungi	decreased	in	grazing	treatments.	 This	conclusion	would	match	results	of	Nadian	et	al.	(1997)	who	also	found	that	soil	compaction	had	no	effect	on	percent	root	colonized,	but	decreased	total	root	colonization	because	root	length	was	greatly	reduced.	Spore	Density		 Unlike	root	colonization,	spore	density	tended	to	be	higher	under	grazing,	at	least	at	the	surface.	This	result	supports	results	of	Frank	et	al	(2003);	however,	studies	have	also	found	decreases	in	spore	density	with	grazing	(Su	and	Guo	2007;	Murray	et	al.	2010),	or	no	changes	at	all	(Yang	et	al.	2013).	The	difference	may	be	associated	with	the	intensity	of	grazing:	Su	and	Guo	(2007)	showed	decreased	sporulation	in	severely	overgrazed	areas	of	the	Mongolian	Steppe	where	plant	richness	and	cover	was	also	reduced.	Similarly,	Ba	et	al	(2012)	found	that	grazing	increased	sporulation,	with	the	largest	increases	at	light	grazing	intensities,	with	less	of	an	increase	at	higher	grazing	intensities.	In	my	study,	grazing	intensity	was	not	intense,	as	I	did	not	detect	any	changes	in	plant	richness,	thus	my	study	supports	studies	showing	increased	sporulation	under	mild	to	moderate	grazing	pressure.	28		The	reasons	for	increases	in	sporulation	associated	with	moderate	grazing	are	not	clear,	but	may	be	a	fungal	response	to	stress.	 For	example,	it	has	been	shown	that	sporulation	is	stimulated	among	the	Glomeromycota	when	fungi	are	carbon	limited	(von	Alten	et	al.	1993).		Presumably	this	is	an	adaptive	strategy	to	improve	survival	(in	the	form	of	resistant	propagules)	during	difficult	times.	Soil	Hyphal	Length		 Soil	hyphal	length	decreased	with	grazing,	more	noticeably	in	the	surface	than	the	subsurface.	 A	reduction	in	SHL	has	been	shown	before	by	some	(Miller	et	al.	1995),	but	not	others	(Eom	et	al.	2001;	Yang	et	al.	2013).	 Soil	hyphal	length	is	not	used	as	often	as	spore	density	and	percent	colonization	to	measure	AM	fungal	infectivity	because	it	is	not	a	discrete	measure	of	AM	fungal	abundance;	other	fungi,	including	saprotrophs	and	parasites	are	included	in	this	measure	(Hart	and	Reader	2002a).	 However,	it	is	considered	a	robust	relative	measure	of	AM	fungal	abundance	in	the	soil	(Newman	1966;	Miller	et	al.	1995;		Hart	and	Reader	2002a).	Reduced	SHL	under	grazing	could	lead	to	reduced	host	tolerance	to	grazing	as	AM		 fungi	are	thought	to	help	plants	recover	from	defoliation	by	increasing	their	nutrient	absorption	(Walling	and	Zabinski	2006)	and	a	reduction	in	hyphal	length	would	theoretically	reduce	nutrient	acquisition	by	the	plant	(Walling	and	Zabinski	2004).		 But	since	I	detected	concurrent	increases	in	N	in	grazed	plots,	reduced	SHL	may	decrease	simply	fungi	may	require	less	hyphae	to	access	the	same	amount	of	nutrients.	Alternatively,	higher	nutrient	levels	in	grazed	plots	may	inhibit	SHL:	 high	levels	of	P	and	N	are	well	known	to	reduce	mycorrhizal	activity	(Treseder	2004;	Jansa	et	al.	2006).	 In	my	study,	although	I	detected	reduced	SHL	and	increased	N	in	grazed	plots,	these	changes	were	not	correlated,	suggesting	that	it	is	another	factor	of	grazing	that	is	causing	the	changes,	not	the	29		related	soil	nutrient	changes.	The	concomitant	changes	in	sporulation	and	hyphal	length	may	have	implications		 for	the	stability	of	the	AM	symbiosis	under	grazing.	Extra-radical	mycelia	are	the	nutrient	acquisition	structures,	and	AM	fungi	with	larger	mycelia	are	thought	to	confer	greater	plant	benefit	(Hart	and	Reader	2002).	Under	grazing	an	extensive	external	mycelium	may	represent	too	much	of	a	carbon	drain	on	hosts	that	are	already	experiencing	carbon	limitation,	particularly	when	nutrients	are	not	limiting	(Hart	and	Reader	2002;	Piippo	et	al.	2011a).	Increased	spore	production	combined	with	the	reduced	nutrient	acquisition	capacity	suggests	that	the	symbiosis	may	not	be	as	beneficial	under	grazing.	In	such	situations,	the	fungi	and	plant	may	act	as	competing	carbon	sinks	(Piippo	et	al.	2011b).	Considering	all	fungal	traits,	it	is	important	to	highlight	that	all	three	variables		 (percent	colonization,	spore	density	and	soil	hyphal	length)	gave	conflicting	measures	of	fungal	abundance.	This	result	speaks	to	the	importance	of	measuring	multiple	measures	of	fungal	abundance.				4.2 Does	grazing	affect	AM	fungal	community	composition?		 Contrary	to	my	expectations,	AM	fungal	communities	did	not	change	in	response	to	grazing.		In	some	studies,	grazing	has	reduced	AM	fungal	diversity	(Ba	et	al.	2012;	Bai	et	al.	2013;	Eom	et	al.	2001;	Su	and	Guo	2007),	caused	shifts	in	relative	abundance	of	AM	fungi	(Eom	et	al.	2001;	Murray	et	al.	2010)	or	had	no	effect	on	the	soil	AM	fungal	community	(Yang	et	al.	2013;	Shelton	et	al.	2014).		 The	lack	of	consensus	is	not	surprising	given	the	multiplicity	of	methods	and	approaches	used	to	quantify	“AM	fungal	communities”.	 In	general,	researchers	use	a	wide	variety	of	approaches	from	spore	counts	to	molecular	inventories	to	obtain	AM	fungal	species	inventories,	making	comparisons	difficult	(Hart	et	30		al.	2016?).		In	general,	studies	looking	at	the	effect	of	disturbance	on	AM	fungal	communities	often	find	only	changes	in	relative	abundance	(Jansa	et	al.	2002;	Jansa	et	al.	2003;	Schnoor	et	al.	2011;	Saito	et	al.	2004;	Murray	et	al.	2010;	Hassan	et	al.	2013;	Sharmah	and	Jha	2014),	rather	than	changes	in	identity.	 In	this	study,	it	is	likely	that	the	co-evolution	of	grassland	AM	fungi	and	grazing	over	millennia	has	made	these	communities	robust	to	moderate	grazing	pressure.	In	this	sense,	Lekberg	et	al	(2012)	suggested	that	the	lack	of	AM	fungal	community	response	to	disturbance	may	be	the	result	of	strong	community	resilience	when	disturbed.	The	fact	that	we	found	no	community	response	to	grazing	in	AM	fungi	may	speak	to		 the	generalist	nature	and/or	plasticity	of	AM	fungi.	If	the	changes	in	AM	fungal	infectivity	were	the	results	of	changes	in	community	composition,	then	it	should	have	been	possible	to	detect	phylogenetic	changes	as	well	–	i.e.	the	community	shifting	from	fungi	that	invest	primarily	in	extra	radical	mycelium	like	Gigasporaceae	(Hart	and	Reader	2002b)	to	those	that	sporulate	more	like	Glomeraceae	(Oehl	et	al.	2009).	Instead,	I	found	that	while	the	relative	abundance	of	AM	fungal	structures	changed,	the	identity	did	not.	 This	may	indicate	that	changes	in	AM	fungal	infectivity	result	from	phenotypic	plasticity	in	response	to	a	change	in	grazing	regime.	Because	AM	fungi	harbour	genetically	different	nuclei	they	are	able	to	rapidly	produce	variable	offspring	with	different	phenotypic	characteristics		(Angelard	et	al.	2014).	This	flexibility	may	allow	them	to	rapidly	respond	to	any	disturbance	without	radical	changes	in	community	composition	Despite	the	lack	of	significant	change	in	overall	community	composition,	some	AM	fungal	isolates	occurred	preferentially	in	grazed	or	ungrazed.	In	my	study,	Glomus	isolates,	plus	Paraglomus	(Paragloemraceae)	and	Ambispora	(Ambisporaceae),	were	indicators	of	grazing.		Not	much	research	has	been	done	on	traits	of	Paraglomeracae	and	31		Ambisporaceae	compared	to	Glomeraceae,	but	the	latter	is	widely	considered	the	more	disturbance	tolerant	AM	fungal	family.	Previous	studies	have	found	that	AM	fungi	in	the	genus	Glomus	are	particularly	tolerant	of	grazing	(Eom	et	al.	2001;	Su	and	Guo	2007;	Ba	et	al.	2012),	perhaps	due	to	their	fast	growth	rates	(Hart	and	Reader	2005)	abundant	and	frequent	sporulation	(Oehl	et	al.	2009).	 While	my	data	support	these	findings,	I	also	show	that,	the	majority	of	Glomeraceae	OTUs	showed	no	preference	between	grazed	or	ungrazed.	I	found	Diversispora	isolates	that	were	indicators	of	ungrazed	plots.	The		 Diversisporaceae	family	is	less	studied	than	the	Glomeraceae,	but	taxa	in	this	family	have	been	found	to	be	both	indicators	of	undisturbed	and	disturbance	sites	(Eom	et	al.	2001;	Moora	et	al.	2014).	Other	members	of	the	order	Diversisporales	are	known	to	have	slower	growth	rates	(Hart	and	Reader	2002b;	Hart	and	Reader	2005)	and	later,	less	frequent	sporulation	(Pringle	and	Bever	2002;	Oehl	et	al.	2009)	than	the	Glomeraceae,	possibly	making	them	less	competitive.		 Unfortunately,	with	the	lack	of	research	into	Diversispora,	traits	of	this	genus	are	relatively	unknown,	making	it	very	difficult	to	determine	why	taxa	in	this	genus	might	be	vulnerable	to	grazing.	4.3 Does	grazing	affect	the	mycorrhizal	environment?	 Plant	and	soil	responses	to	grazing	AM	fungal	grazing	responses	were	unrelated	to	environmental	responses	to	grazing	even	though	grazing	induced	several	changes	to	plant	and	soil	characteristics.	Plant	community		 In	agreement	with	my	hypothesis,	grazing	reduced	plant	biomass	and	changed	plant	evenness,	although	plant	richness	was	not	affected.	These	findings	support	other	studies	that	found	no	changes	in	species	richness	with	grazing,	especially	in	dry	and	nutrient	poor	32		sites	(Lkhagva	et	al.	2013;	Frank	2005).	Like	other	studies	before,	plant	evenness	increased	with	grazing	(Frank	2005;	Ba	et	al.	2012;	Krzic	et	al.	2012).	This	may	be	explained	by	trade	offs	among	plant	taxa	in	competitive	ability	and	grazing	tolerance	(van	Velzen	and	Etienne	2015).	For	example,	in	our	study,	ungrazed	plots	were	dominated	by	Festuca,	which	are	highly	palatable,	but	grazing	intolerant	(Mack	and	Thompson	1982).	They	are	well	known	as	a	‘decreasers’	(Wikeem	et	al.	2012;	van	Ryswyk	et	al.	1966)	and	display	several	traits,	such	as	height	and	tussock	architecture,	that	are	not	favoured	by	grazing	(Diaz	et	al.	2007).	In	the	absence	of	grazing,	this	plant	was	able	to	outcompete	many	others	and	dominate	the	exclosures,	resulting	in	a	decrease	in	plant	evenness	in	ungrazed	plots.	Are	plant	community	changes	responsible	for	changes	in	AM	fungal		infectivity/community?		 That	changes	in	plant	community	was	not	related	to	changes	in	AM	fungal	community	is	surprising,	as	other	grazing	studies	have	linked	changes	in	AM	fungal	community	with	grazing	to	plant	richness	(Yang	et	al.	2013),	and	plant	community	(Valyi	et	al.	2015)	responses.	 In	fact,	plant	community	composition	is	generally	the	most	important	driver	for	changes	in	AM	fungal	communities	(Davison	et	al.	2011;	Opik	et	al.	2014).	 This	is	largely	believed	to	result	from	preference	among	AM	taxa	and	their	hosts	(Opik	et	al.	2014;	Torrecillas	et	al.	2012).	In	our	study,	changes	in	plant	community	were	due	to	changes	in	eveness,	not	species	turnover.	 Further,	this	was	largely	due	to	one	species	(Festuca).	 Thus,	preferred	hosts	were	still	available	for	AM	fungi,	even	under	grazing.	 Additionally,	since	plants	typically	host	multiple	fungi,	and	fungal	isolates	are	connected	to	multiple	plants	(Vandenkoornhuyse	et	al.	2003),	the	fungi	associated	with	Festuca	in	grazed	plots	were	likely	already	well	adapted	to	other,	co-occuring	taxa..	Soil	Nutrients	33			 As	predicted,	grazing	increased	soil	nutrients		 (available	P	and	total	N,	minimally	affected	soil	carbon).	The	increase	in	soil	N	with	grazing	is	consistent	with	the	literature	(Wallace	1987;	Seagle	et	al.	1992;	Shariff	et	al.	1994;	Frank	and	Evans	1997).	Increased	soil	N	may	be	explained	by	the	return	of	N	through	defecation	and	urine	(Frank	and	Evans	1997;	Holland	et	al.	1992)	as	well	as	feedbacks	between	plant	responses	to	grazing	and	N	cycling	(Holland	et	al.	1992;	Frank	and	Groffman	1998;	Hamilton	et	al.	2008).	Grazing	causes	an	increase	in	N	mineralization,	which	under	grazing	can	be	double	that	of	ungrazed	plots	(Frank	and	Groffman	1998).	Hamilton	et	al	(2008)	demonstrated	that	defoliation	increased	carbon	exudation	in	the	roots,	which	led	to	increased	N-mineralization	in	the	rhizosphere	by	stimulating	soil	microbes.	This	link	between	carbon	exudation	and	N-	mineralization	may	explain	why	total	C	and	N	were	so	highly	correlated.	While	there	are	many	studies	examining	the	effects	of	grazing	on	the	N	cycle	(Holland	and	Detling	1990;	Frank	and	Evans	1997;	Olofsson	et	al.	2001)	available	P	responses	seem	understudied.	Likely,	this	is	because	most	grasslands	are	N-limited		(Vitousek	and	Howarth	1991)	making	P	availability	relatively	unimportant.	Of	the	few	that	have	examined	the	response	of	available	P	to	grazing,	one	found	a	decrease	(Xie	and	Wittig	2004),	and	another	found	no	change	(Baron	et	al.	2001),	unlike	my	results	which	indicate	a	weak	trend	for	increased	P	levels	under	grazing.	Soil	compaction		 As	predicted,	I	found	that	soils	were	compacted	by	grazing.	My	results	agree	with	others	who	found	higher	bulk	density	under	grazing	down	to	15cm	(Krzic	et	al.	2012).	In	the	deeper	soils	I	found	no	significant	differences	between	grazed	and	ungrazed,	most	likely	because	compaction	affects	the	surface	layers	most	(Xie	and	Wittig	2004;	Krzic	et	al.	34		2012).	Soil	compaction	is	a	well-known	consequence	of	grazing	(Chanasyk	and	Naeth	1995;	Greenwood	and	McKenzie	2001;	Xie	and	Wittig	2004),	though	few	studies	focused	on	microbial	communities	choose	to	measure	it.	Are	changes	in	soil	environment	causing	changes	in	AM	fungal	infectivity/community?		 Despite	the	significant	effect	of	grazing	on	soil	properties,	none	of	these	factors	were	related	to	AM	fungal	grazing	responses.	Other	studies	have	linked	grazing	induced	changes	in	AM	fungal	community	composition	to	soil	fertility	(Murray	et	al.	2010;	Yang	et	al.	2013),	or	soil	electrical	conductivity	(Ba	et	al.	2012);	but	in	my	study	soil	properties	had	no	effect.	This	is	supported	by	Shelton	et	al	(2014),	who	also	found	no	difference	in	AM	fungal	communities	despite	significant	changes	in	soil	density.	In	addition,	the	soil	bulk	density	under	grazing	(0.818	g/cc),	though	higher	than	the	average	bulk	density	in	ungrazed	(0.676g/cc)	does	not	reach	levels	(~1.4g/cc)	known	to	impede	root	growth	(Atwell	1993).	Ultimately,	that	AM	fungal	responses	were	unrelated	to	plant	and	soil	changes	indicates	that	1)	AM	fungi	are	resistant/tolerant	of	these	changes,	and/or	2)	AM	fungal	changes	are	more	related	to	environmental	variables	I	did	not	measure	(i.e.	root	biomass).	1) AM	fungi	resist/tolerate	changes	to	soil	environment	supports	the	idea	that	the	AM	fungi	are	generalist	and	have	high	phenotypic	plasticity	(Angelard	et	al	2014).	This	would	explain	why	the	community	composition	did	not	change	significantly	–	community	members	were	able	to	adapt	to	new	conditions.	 The	genotypic	and	phenotypic	flexibility	of	AM	fungi	means	that	isolates	would	be	able	to	shift	functioning	as	necessary	to	survive	the	pressure	of	grazing	in	an	area	(Angelard	et	al.	2014).	This	may	be	related	to	high	levels	intra-isolate	variation	found	in	AM	fungi	(Munkvold	et	al.	2004),	but	more	research	is	needed	to	understand	the	link	between	genetic	and	trait	plasticity	in	AM	fungi.	2) AM	fungi	are	driven	by	unaccounted	for	factors	That	there	were	significant	changes	in	35		spore	density	and	hyphal	length	to	grazing	means	that	AM	fungi	were	affected	by	something,	but	it	may	not	be	plant	community	and	the	soil	properties	measured	in	my	study.	One	such	variable	may	be	root	length.	Reduced	root	length	means	less	total	root	colonization,	which	may	explain	the	decrease	in	soil	hyphal	length	with	grazing,	since	smaller	root	mycelium	may	mean	fewer	resources	for	external	mycelium	production.	Decreased	root	availability	may	also	lead	to	increased	spore	abundance	because	spores	can	stay	dormant	if	there	is	no	root	to	colonize	(Smith	and	Read	2008).	This	is	one	possible	variable	that	was	not	measured	but	may	help	explain	the	AM	fungal	responses	to	grazing.		4.4 The	effect	of	time	As	predicted,	the	age	of	the	exclosure	was	the	most	important	factor	explaining	grazing	related	changes	in	AM	fungal	community	composition.	 That	this	was	not	true	for	measures	of	AM	fungal	infectivity	implies	that	AM	fungal	infectivity	responses	to	grazing	happen	relatively	quickly;	quicker	than	changes	to	the	plant	communities,	which	may	take	upwards	of	50	years	to	recover	from	grazing	(Pyke	et	al.	2016;	Krzic	et	al.	2012).	The	relationship	between	age	of	exclosure	and	AM	fungal	dissimilarity	suggest	a	succession-like	change	over	time.	In	order	to	show	true	evidence	of	succession,	I	would	need	to	show	that	the	AM	fungal	communities	were	moving	in	a	certain	direction,	which	was	not	possible	with	my	data	set.	The	change	in	fungal	communities	has	been	a	topic	of	interest	for	several	decades	(Janos	1980;	Johnson	et	al.	1991;	Boerner,	DeMars	&	Leicht	1996;	Treseder	et	al.	2004;	Davison	et	al.	2011),	but	most	studies	are	limited	to	short	time	scales	or	conflate	site	with	time.	In	our	study,	the	paired	design	allowed	us	to	compare	grazing	effects	among	communities	of	the	same	age.	The	fact	that	the	age	of	the	exclosure	is	the	only	factor	related	to	fungal	community	changes,	and	the	dissimilarity	increases	with	age	underlines	the	36		importance	of	long-term	studies	(over	six	decades	in	this	case).	37		CHAPTER	5:	CONCLUSION					 My	goal	was	to	determine	how	grazing	affect	AM	fungi	in	temperate	grasslands,	and	with	this	study	I	was	able	to	conclude	that	grazing	affects	AM	fungal	infectivity	more	than	it	changes	the	community	composition.	 Further,	I	provided	evidence	of	long	term	differentiation	in	AM	fungal	communities,	in	that	grazed	plots	become	more	dissimilar	from	ungrazed	plots	until	over	time.				5.1 AM	fungal	infectivity		 I	showed	that	discovering	the	changes	in	AM	fungal	infectivity	requires	multiple	measures,	and	reliance	on	percent	colonization	in	the	literature	may	cause	researchers	to	miss	important	changes.		These	changes	may	also	have	functional	implications;	the	decrease	in	hyphal	length	and	increase	in	spore	density	may	be	detrimental	to	the	plant	but	this	hypothesis	requires	further	study.	Unfortunately,	I	was	not	able	to	link	the	changes	in	AM	fungal	infectivity	with	any	environmental	changes,	which	is	most	likely	because	other	factors	may	be	driving	the	differences	in	infectivity	between	grazed	and	ungrazed	plots	(i.e.	root	biomass).				5.2 AM	fungal	community		 The	AM	fungal	community	composition	was	surprisingly	resilient	to	grazing	pressure.	Considering	the	changes	in	AM	fungal	infectivity,	I	had	assumed	the	community	would	shift	from	AM	taxa	with	high	levels	of	SHL	(i.e.	Gigasporaceae)	to	those	taxa	known	to	be	prolific	sporulaters	(i.e.	Glomeraceae).	That	this	taxonomic	shift	does	not	occur	indicates	the	AM	fungal	communities	maybe	have	high	trait	plasticity,	and	are	able	to	respond	to	grazing	38		without	turnover	of	taxa.	This	idea	is	reinforced	by	the	lack	of	relationship	between	changes	in	plant	and	soil	characteristics	and	AM	fungal	community	composition.				5.3 The	Effect	of	Time		 I	show	robust	evidence	that	time	plays	a	role	in	the	change	of	AM	fungal	communities	with	grazing.	I	found	a	strong	increase	in	AM	fungal	community	dissimilarity	with	time;	meaning	communities	inside	and	outside	the	exclosures	became	more	dissimilar	the	longer	the	exclosure	has	been	in	place.	 This	highlights	the	importance	of	long-term	studies	in	order	to	understand	the	relationship	between	grazing	and	AM	fungal	communities.	39		FIGURES							 Figure	1:	The	standardized	change	in	AM	fungal	infectivity	with	grazing.	Change	=	(infectivity	in	grazed)	–	(infectivity	in	ungrazed).	The	solid	red	line	indicates	no	difference	between	grazed	and	ungrazed.	Open	circles	show	the	change	at	each	site,	closed	circles	represent	the	standardized	median,	and	error	bars	show	the	95%	confidence	intervals.	Col	=	Colonization,	Hyph	=	Hyphal	length,	Spore	=	Spore	density,	Sur	=	surface	soil,	Sub	=	subsurface	soil.	 P-values	of	the	paired	Wilcox	signed	rank	test	were	as	follows:	Col	Sur	(p-	value=0.930),	Hyph	Sub	(p-value	=	0.160),	Hyph	Sur	(p-value	=	0.039),	Spore	Sub	(p-	value=0.359),	Spore	Sur(p-value=0.055).	N=9	40			 Figure	2:	Nonmetric	multidimensional	scaling	(NMDS)	showing	distance	between	AM	fungal	communities	from	grazed	and	ungrazed	plots	for	both	soil	depths	(data	pooled).	Ordination	axes	were	calculated	using	Bray-Curtis	distance	metric	(Bray	and	Curtis	1957),	with	4	axes	and	a	stress	value	of	0.1746.	 Grazing	was	not	a	significant	factor	any	of	the	axes.	An	ordination	of	each	layer	separately	also	finds	no	significant	effect	of	grazing	on	any	of	the	axes.	40			 Figure	3:	The	standardized	change	in	AM	fungal	richness	and	evenness	with	grazing.	The	solid	red	line	indicates	no	difference	between	grazed	and	ungrazed.	Open	circles	show	the	change	at	each	site,	closed	circles	represent	the	standardized	mean,	and	error	bars	show	the	95%	confidence	intervals.	Sur	=	surface	soil,	Sub	=	subsurface	soil.	P-values	for	the	paired	Wilcox	signed	rank	test	were	as	follows:	Evenness	Sub	(p-value=0.301),	Evenness	Sur	(p-value	=	0.735),	Richness	Sub	(p-value	=	0.875),	Richness	Sur	(p-value	=	0.938).	N=9	41		A	 B																														Figure	4:	The	relationship	between	Age	of	the	exclosures	and	the	dissimilarity	between	AM	fungal	communities	in	grazed	and	ungrazed.	Plot	A	represents	the	surface	communities	and	plot	B	shows	the	subsurface	communities.	Surface	r=0.66,	subsurface	r=0.64.	42			 Depth	 Effect	size	 95%	CI	 P	value	 Adjusted2	P	value	Plant	Community	 	 	 	 	 	Plant	Evenness	 	 0.145	 0.005,	0.258	 0.039	 0.117	Plant	Richness	 	 3.25	 -3.5,	9.5	 0.414	 1.000	Plant	Biomass	 	 -36.2	 -61.6,	-24.1	 0.004	 0.012	Soil	Properties	 	 	 	 	 	Total	C	 Surface	 0.535	 -0.24,	1.18	 0.0742	 0.370	(mg/kg)	 Subsurface	 -0.045	 -0.42,	0.27	 0.8203	 1.000	Total	N	 Surface	 0.056	 0.007,	0.106	 0.039	 0.195	(mg/kg)	 Subsurface	 -0008	 -0.033,	0.0185	 0.5703	 1.000	P	 Surface	 4.465	 -1.770,	28.765	 0.0742	 0.370	(mg/kg)	 Subsurface	 1.305	 -0.07,	35.79	 0.0547	 0.273	pH	 Surface	 0	 -0.19,	0.155	 0.992	 1.000		 Subsurface	 -0.0225	 -0.38,	0.35	 0.6406	 1.000	Bulk	Density1	 Surface	 0.133	 0.0935,	0.205	 0.0078	 0.039	(g/cc)	 Subsurface	 0.0347	 -0.0268,	0.0936	 0.1953	 1.000		TABLES		Table	1:	Description	of	the	nine	exclosures	used	in	this	study.	Elevation	  Site Latitude	(N) Longitude	(W) (m) Slope Age*   Muscrat	Lk	 50°	8'2.15"	 120°26'30.45"	 1025	 30	 17	Tunkwa	 New	 50°35'53.45"	 120°51'55.15"	 1150	 4	 21	Bluegrass	 50°	4'45.27"	 120°28'2.70"	 1000	 0	 21	Aspen	 50°	4'54.14"	 120°25'55.74"	 1200	 7	 37	Stipa	rich	 50°	3'58.90"	 120°26'51.29"	 1125	 8	 42	Stipa	nel	 50°	4'43.39"	 120°26'58.88"	 1000	 5	 48	Tunkwa	Old	 50°35'58.03"	 120°51'57.96"	 1150	 0	 55	Repeater	 50°	4'32.08"	 120°25'30.02"	 1306	 0	 65	Goose	Lk	 50°	6'11.59"	 120°27'55.19"	 1160	 5	 85	*Age	indicates	age	of	exclosure		Table	2:	Biotic	and	abiotic	responses	to	grazing	at	nine	grazing	exclosures.	Differences	 in	plant	and	soil	variables	between	grazed	and	ungrazed	were	tested	using	Wilcoxon	signed	rank	tests.	95%	confidence	intervals	than	did	not	overlap	with	0	and	p-values	less	than	0.1	  are	bolded	as	an	indication	that	there	was	some	difference	between	grazed	and	ungrazed.																					 1	Bulk	densities	were	collected	from	8	sites	only	because	one	of	the	sites	was	too	rocky	2Adjusted	P	values	were	calculated	using	the	bonferroni	correction	for	multiple	comparisons.	43		Table	3:	The	change	in	percent	cover	with	grazing	of	the	most	abundant	plant	species.	Only	plants	that	were	found	in	most	sites	(at	least	7/9)	and	made	up	a	significant	proportion	(at	least	10%)	of	the	plant	community	were	included.	Change	was	calculated	as	=	(%	cover	in	grazed)-(%cover	in	ungrazed)     Plant G* A B S T O H M R   Bromus	inermis	 0	 0	 1	 0	 -5.1	 -11.55	 0.3	 0	 0	Elymus	spicata	 12	 0	 0	 0.35	 -0.6	 0	 0	 37.75	 0	Festuca	campestris	 -89.3	 -36.4	 0	 -38.5	 -26.05	 -62.5	 -1.05	 -55.3	 -75.4	Koeleria		macrantha	 4.8	 13.2	 0	 -0.15	 5.25	 5.95	 0	 -1	 1.9	Phleum	pratense	 0	 0	 13.55	 0	 0	 0	 0	 0	 0	Poa	pratensis	 -4.3	 18.45	 -3.2	 34.25	 -14.7	 20.8	 18.6	 0	 6.5	Poa	secunda	 16.2	 1.25	 0	 0	 3.3	 -12.1	 0	 5.2	 4.1	Stipa	nelsonii	 7.3	 1.25	 -0.05	 10.55	 -4.4	 -1.05	 -0.7	 0	 0.8	Stipa	richardsonii	 0	 0	 0	 0.45	 14.8	 22.6	 6.75	 0.05	 0.2	Achillea		millefolium	 -8.9	 -3.1	 1.5	 6.5	 3.1	 12.7	 0.9	 0.3	 -1	Agroseris	glauca	 -13.2	 -0.1	 -0.65	 -3.05	 0	 0	 -0.65	 0	 -4.4	Antennaria	dimorpha	 	 2.5	 	 -1.15	 	 0	 	 0	 	 -2.95	 	 0.05	 	 0	 	 0.55	 	 0.1	Antennaria	 	 	 	 	 	 	 	 	 	microphylla	 2.2	 0	 0	 0	 1.95	 2	 0	 3.6	 16.9	Aster	campestris	 0.5	 2.65	 -0.4	 -0.35	 -0.9	 2.85	 0.55	 -0.05	 -1.1	Astragalus	miser	 0.1	 0.3	 -0.3	 1.25	 0	 -0.7	 -0.1	 0.15	 1.9	Cerastium	arvense	 0	 -1	 0.9	 -0.1	 9.6	 17.5	 0.4	 0	 0	Fragaria		virginiana	 -9.5	 0	 0	 0	 -0.1	 -0.4	 0.3	 0	 0	Geum	triflorum	Taraxacum	 0	 -0.35	 -0.15	 -2.25	 -1.3	 0	 -0.45	 0.05	 -0.8	officinale	 -0.1	 -0.2	 25.9	 0	 1.45	 2.1	 0.05	 0.45	 0.1	Tragopogon	dubius	 0	 0	 0.35	 0.25	 0.65	 0.7	 0.25	 0.6	 0.1	*	Site	names	are	abbreviated	as	follows:	G	for	Goose,	A	for	Aspen,	B	for	Bluegrass,	S	for	Stipanel,	T	for	Tunkwa	New,	O	for	Tunkwa	Old,	H	for	Stiparich,	M	for	Muscrat	and	R	for	Repeater.	44		AM	Family1	 Grazed	 Ungrazed	 Stat	 	 P	value	Archeaosporaceae.4	 0	 1	 	 0.4401	 0.9165	Acaulosporaceae.3	 0	 1	 	 0.2883	 0.4312	Diversisporaceae.8	 0	 1	 	 0.7112	 0.1791	Claroideoglomeraceae.8	 0	 1	 	 0.7017	 0.8275	Glomeraceae.33	 1	 0	 	 0.7080	 0.7954		Table	4:	Blocked	indicator	species	analysis	of	AM	fungal	Families	and	isolates	as	indicators	of	grazed	or	ungrazed	plots.	Only	Families	with	more	than	2	isolates	were	included	in	the	family	analysis.	Only	the	nine	strongest	indicator	isolates	were	shown.   									  Paraglomeraceae.3 1 0 0.6062 0.1342   AM	OTU	Glomus	VTX00342	 1	 0	 0.5586	 0.0031	Diversispora	VTX00054	Paraglomus	brasilianum	VTX00239	0		1	1		0	0.5937		0.4419	0.0388		0.0817	Ambispora	leptoticha	VTX00242		1		0		0.2934		0.084	Glomus	VTX00144	 1	 0	 0.3375	 0.0856	Diversispora	VTX00062	 0	 1	 0.7142	 0.0974	DiversisporaVTX00347	Septoglomus	furcatum	VTX00064	0		1	1		0	0.4591		0.7174	0.1024		0.1251	Glomus	VTX00172	 0	 1	 0.5457	 0.1392	1	Numbers	after	each	fungal	family	indicate	how	many	OTUs	belonged	to	each	family	45		Table	5:	Summary	of	Multimodel	Inference	Statistics.	Italicized	descriptions	are	the	dependent	values	for	the	global	models.	Model	summaries	are	in	the	Appendix			Variable	 Stdized	Estimate	 	 Std.	Error	 	 95%	CI	 Relative	Importance		Difference	in	Soil	Hyphal	Length	between	grazed	and	ungrazed	in	the	surface	soil		 ΔN	 0.118	 0.265	 -0.468,	1.220	 0.31	ΔP	 0.078	 0.229	 -0.567,	1.187	 0.25	ΔPlant	evenness	 0.012	 0.164	 -0.865,	1.007	 0.17	Age	 0.012	 0.159	 -0.843,	0.983	 0.17	Plant	dissimarity	 -0.009	 0.16	 -0.979,	0.874	 0.16		Difference	in	Spore	Density	between	grazed	and	ungrazed	in	the	surface	soil		 ΔPlant	evenness	 0.313	 0.381	 -0.196,	1.375	 0.53	ΔP	 0.083	 0.231	 -0.454,	1.24	 0.21	Plant	dissimilarity	 0.011	 0.176	 -1.005,	1.167	 0.13	ΔN	 0.02	 0.137	 -0.713,	1.065	 0.11	Age	 -0.011	 0.127	 -1.006,	0.792	 0.11	AM	fungal	dissimilarity	between	grazed	and	ungrazed	in	the	surface	soil	Age	 0.526	 0.391	 0.099,	1.344	 0.73	ΔN	 0.194	 0.299	 -0.088,	1.144	 0.37	Plant	dissimilarity	 0.06	 0.203	 -0.345,	1.314	 0.12	ΔP	 -0.057	 0.205	 -1.375,	0.388	 0.12	ΔPlant	evenness	 0.006	 0.103	 -0.898,	1.150	 0.05	AM	fungal	dissimilarity	between	grazed	and	ungrazed	in	the	subsurface	soil	Age	 0.604	 0.379	 0.095,	1.386	 0.81	ΔPlant	evenness	 -0.108	 0.233	 -1.090,	0.168	 0.23	ΔN	 -0.040	 0.164	 -1.187,	0.437	 0.11	Plant	dissimilarity	 -0.051	 0.180	 -1.226,	0.340	 0.11	ΔP	 0.026	 0.128	 -0.443,	1.078	 0.08	46		BIBLIOGRAPHY		Aguilar-Chama	A.,	and	Guevara	R.	2012.	“Mycorrhizal	Colonization	Does	Not	Affect	Tolerance	to	Defoliation	of	an	Annual	Herb	in	Different	Light	Availability	and	Soil	Fertility	Treatments	but	Increases	Flower	Size	in	Light-Rich	Environments.”	Oecologia	168	(1):	131–39.	doi:10.1007/s00442-011-2066-1.		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Journal	of	Vegetation	Science	1-8.	doi:10.1111/jvs.12191.	58		APPENDIX	A:	SUPPLEMENTARY	MATERIALS						Ungrazed				 Archeosporaceae	Acaulosporaceae	Diversisporaceae	Claroideoglomeraceae	Paraglomeraceae	Glomeraceae					Grazed				 Archeosporaceae	Acaulosporaceae	Diversisporaceae	Claroideoglomeraceae	Paraglomeraceae	Glomeraceae				Figure	5:	Abundance	of	AM	fungal	families	in	grazed	and	ungrazed	plots.	All	sites	are	pooled	together	for	this	figure.	59		2	 3	 -12.57	 35.95	 0	 0.28	1	 3	 -12.81	 36.42	 0.47	 0.22	4	 3	 -13.23	 37.27	 1.32	 0.15	5	 3	 -13.25	 37.29	 1.35	 0.14	3	 3	 -13.25	 37.31	 1.36	 0.14	12	 4	 -12.46	 42.92	 6.97	 0.01	25	 4	 -12.47	 42.95	 7	 0.01	23	 4	 -12.53	 43.05	 7.11	 0.01	24	 4	 -12.57	 43.14	 7.19	 0.01	15	 4	 -12.6	 43.2	 7.25	 0.01	14	 4	 -12.74	 43.47	 7.52	 0.01	13	 4	 -12.76	 43.52	 7.57	 0.01	34	 4	 -13.13	 44.25	 8.31	 0	35	 4	 -13.22	 44.44	 8.49	 0		Statistical	Models	in	R	AM	fungal	Community	comm.ord	=	metaMDS(comm.t[,5:ncol(comm.t)],	k	=	4,	trymax	=	50)		 axis1.mod	=	lmer(NMDS1~Description+	(1|Site:Description)+	(1|Site),	data	=	comm.char)	axis2.mod	=	lmer(NMDS2~Description+	(1|Site:Description)+	(1|Site),	data	=	comm.char)	axis3.mod	=	lmer(NMDS3~Description+	(1|Site:Description)+	(1|Site),	data	=	comm.char)	axis4.mod	=	lmer(NMDS4~Description+	(1|Site:Description)+	(1|Site),	data	=	comm.char)	Multimodel	inference	Models	Parameter	codes:	1=Age,	2=	Change	in	N,	3	=	Change	in	P,	4=	Plant	dissimilarity,	5=Change	in	plant	evenness		 Global	Model:	AMF.test	=	lm(difference	in	AMF~	Age+	change	in	N	+	change	in	P	 +	plant.diss	+	change	in	plant	evenness,	data	=	data,	na.action	=	na.fail)		Table	6:	Summary	of	Model	Selection	Statistics	for	the	Surface	Soil	Change	in	Hyphal	Length.	Models	showing	ΔAICc	less	than	10	are	shown	here.	  Model				df logLik AICc ΔAICc				weight   																					  45 4 -13.22				 44.45 8.5 0   	Table	7:	Summary	of	Model	Selection	Statistics	for	the	Surface	Soil	Change	in	Spore	Density.	Models	showing	ΔAICc	less	than	10	are	shown	here.	60			Model	 df	 logLik	 AICc	 ΔAICc	 weight	4	 3	 -55.5	 121.79	 0	 0.46	1	 3	 -56.44	 123.67	 1.88	 0.18	3	 3	 -57.07	 124.94	 3.14	 0.1	2	 3	 -57.1	 125	 3.2	 0.09	5	 3	 -57.25	 125.3	 3.51	 0.08	34	 4	 -54.77	 127.54	 5.75	 0.03	45	 4	 -55.21	 128.42	 6.62	 0.02	14	 4	 -55.43	 128.85	 7.06	 0.01	24	 4	 -55.49	 128.97	 7.18	 0.01	13	 4	 -56.32	 130.64	 8.85	 0.01	15	 4	 -56.4	 130.79	 9	 0.01	12	 4	 -56.44	 130.87	 9.08	 0	  35 4 -56.82				 131.65				9.85 0   			Table	8:	Summary	of	Model	Selection	Statistics	for	the	AM	fungal	dissimilarity	in	the	Surface	Soil.	Models	showing	ΔAICc	less	than	10	are	shown	here.		Model	 df		 logLik	 AICc	 delta	 weight	1	 	 3	 18.1	 -25.39	 0	 0.43	12	 	 4	 21.11	 -24.21	 1.18	 0.24	4	 	 3	 16.82	 -22.85	 2.55	 0.12	2	 	 3	 16.23	 -21.66	 3.73	 0.07	3	 	 3	 15.79	 -20.78	 4.61	 0.04	5	 	 3	 15.66	 -20.53	 4.86	 0.04	14	 	 4	 18.36	 -18.71	 6.68	 0.02	23	 	 4	 18.3	 -18.61	 6.78	 0.01	13	 	 4	 18.14	 -18.28	 7.11	 0.01	15	 	 4	 18.1	 -18.19	 7.2	 0.01	45	 	 4	 17.47	 -16.94	 8.46	 0.01	34	 	 4	 17.47	 -16.93	 8.46	 0.01	  24 4 17.44 -16.87 8.52 0.01   	Table	9:	Summary	of	Model	Selection	Statistics	for	the	AM	fungal	dissimilarity	in	the	Subsurface	Soil.	Models	showing	ΔAICc	less	than	10	are	shown	here.		Model	 df	 logLik	 AICc	 ΔAICc	 weight	5	 3	 16.24	 -21.68	 0	 0.47	45	 4	 18.9	 -19.8	 1.88	 0.18	35	 4	 18.1	 -18.2	 3.48	 0.08	2	 3	 14.5	 -18.2	 3.48	 0.08	15	 4	 17.69	 -17.39	 4.29	 0.05	  4 3 13.86 -16.92 4.76 0.04   61			1	 3	 13.43	 -16.05	 5.63	 0.03	25	 4	 16.93	 -15.87	 5.81	 0.03	3	 3	 13.31	 -15.81	 5.87	 0.02	  24 4 15.61 -13.22 8.47 0.01   

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