The Role Of Endohyphal Symbionts In Influencing Fungal Degradation Of Plant Cell Walls Under Field Conditions
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Authors Thomas, Kendra Allyson
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THE ROLE OF ENDOHYPHAL SYMBIONTS IN INFLUENCING FUNGAL
DEGRADATION OF PLANT CELL WALLS UNDER FIELD CONDITIONS
By
KENDRA ALLYSON THOMAS
______
A Thesis Submitted to The Honors College
In Partial Fulfillment of the Bachelors degree With Honors in
Microbiology
THE UNIVERSITY OF ARIZONA
M A Y 2 0 1 9
Approved by:
______
Dr. A. Elizabeth Arnold School of Plant Sciences
1 The role of endohyphal symbionts in influencing fungal degradation of plant cell walls
under field conditions
Abstract
Many plant-associated fungi form symbiotic relationships with endohyphal bacteria (EHB), which live within fungal tissue. Our goal was to evaluate how EHB (here, Pantoea spp.) influence the ability of fungi to degrade plant material under field conditions, thus extending previous research based only on in vitro assays. We examined the degradation ability of two fungi (9133 and 9140) with (+) or without (-) EHB on foliage of three plant species (Platycladus orientalis, Cupressus arizonica, Juniperus deppeana). Plant material with one treatment (9133+, 9133-, 9140+, 9140-, or control) was placed into mesh litter bags and deployed into field plots. After three months we found that treatment with fungi, regardless of EHB status, increased mass loss relative to controls for two species (C. arizonica, J. deppeana). In contrast to expectations based on in vitro results and a small pilot study, EHB did not increase degradation of plant material by fungi. We anticipate that biotic interactions with other decomposers in soil, especially in a relatively rainy winter with robust microbial activity, could explain the observed results. Future experiments might focus on how EHB influence interactions among the diverse fungi that play a role in plant tissue degradation in complex natural systems.
Introduction
Diverse plant-associated fungi form symbiotic relationships with endohyphal bacteria (EHB), which live within fungal tissue (Shaffer et al. 2017). Many EHB influence the phenotype of their host fungi in ways that can increase the ability of fungi to survive and proliferate in certain environments (Baltrus et al.
2017). For example, EHB can control expression of proteins or hormones by plant-symbiotic fungi that benefit both partners in planta (Hoffman et al. 2013).
Diverse plant-associated and plant-symbiotic fungi have saprotrophic life phases, during which they live outside healthy plants and degrade plant tissue (Blanchette 1991). Recent findings indicate that some EHB can influence the ability of fungi to express enzymes necessary for degradation of cellulose and lignin, the primary constituents of plant cell walls (Arendt 2015, Araldi-Brondolo et al. 2017). This
2 discovery is important because these enzymes are of great interest in bioremediation, biofuel development using plants and plant products, and recycling (Novotny et́ al., 2004, Wu et al. 2017). At this time, observations that EHB can influence fungal degradation of plant cell walls have been confined to in vitro experiments (Arendt 2015, Woytenko et al. in preparation). It is not yet known whether EHB can influence fungal degradation of plant material under natural conditions.
The goal of this thesis was to quantify the influence of EHB on fungal degradation of plant material under field conditions. We focused on two plant-associated fungi, strains 9133 and 9140.
Previous work has shown that 9133 and 9140 express cellulase activity in vitro (Woytenko et al. in preparation). Moreover, the ability of these fungi to degrade plant material in vitro is altered by the presence of EHB (Woytenko et al. in preparation). Both fungi were isolated originally from foliage of
Cupressaceae, a family of conifers that includes junipers and cypresses. Leaf material of conifers often is recalcitrant to degradation because it is tough and rich in secondary metabolites (Neale et al. 2004). The dominant trees in montane ecosystems across the southwestern United States are conifers, and degradation of their leaves and tissues is important in soil production, function, and fertility (Alfredsson et al. 1998). Understanding how EHB influence fungal breakdown of conifer leaf material in a natural environment may provide a new tool to enhance soil quality for reforestation and after fire events.
Here we examined how EHB impact the degradation of conifer foliage by fungal strains 9133 and
9140 under field conditions. We inoculated leaf litter in soil-permeable litter bags and measured mass loss after a three-month incubation period in winter 2018-2019. We predicted that mass loss would vary among the plant species and as a function of fungal treatment, and that EHB would influence mass loss in a manner consistent with their previously observed impacts on plant tissue in vitro. Specifically, we anticipated a positive effect of EHB based on a pilot project and in vitro trials, which indicated that EHB were associated with increased tissue degradation and mass loss relative to the same fungi without EHB.
Materials and Methods
Litter bag construction
We prepared 300 litter bags from 50 m mesh cloth (Kissell et al. unpublished). Each bag was approximately 5 x 10 cm in size. The bags were impervious to insects but porous to water, soil particles,
3 and microbes. The bags were wrapped in aluminum foil and autoclaved on the Gravity 30 cycle before plant material was placed inside.
Collection of plant materials
We collected sensesced leaf material from three species of Cupressaceae in early fall 2018: Platycladus orientalis, Cupressus arizonica, and Juniperus deppeana. Material from P. orientalis and C. arizonica was collected at the University of Arizona Campus Arboretum, primarily from individuals located near Old
Main. The University of Arizona campus is located in urban Tucson at 3213’54.8”N, -11057’12.5”W and
728m above sea level (m.a.s.l.). Both P. orientalis and C. arizonica were planted on campus, but the latter species is native to mountains in southern Arizona. In turn, P. orientalis is native to Asia. We collected material from at least two individuals per each of these species. We collected leaf material from five individuals of J. deppeana on the hillslopes immediately east of the Molino Basin Campground
(coordinates 3220’09.0”N, 11041’57.3”W, 1370 m.a.s.l.), where it occurs natively. We washed the leaf material gently in distilled water and then autoclaved it for one hour on a Gravity 30 cycle prior to placement in the litter bags and inoculation, as described below.
Preparation of inoculum
We initially selected three fungal strains (9133, 9140, and 9143) for the experiment. Each strain was prepared to contain EHB (EHB+), which was the original state in which the fungi were isolated, and to lack EHB (EHB-), achieved by cultivating the fungal strains on antibiotic medium (see below).
To initiate growth of each strain we plated samples from vouchers onto 100 mm agar plates. The strains containing EHB (9133+, 9140+, and 9143+) were plated on 2% MEA (AMRESCO). The strains cured of their EHB (9133-, 9140-, 9143-) were plated on 2% MEA (AMRESCO)+KCAT. The KCAT medium contained 100g/L of ampicillin, 15g/L of tetracycline, 50g/L of kanamycin, and 40g/L of ciprofloxacin (Arendt et al. 2016). Each fungal species with/without bacteria was plated three times.
After ca. two weeks, we extracted total genomic DNA from each culture with the Sigma Extract-n-
Amp Plant Kit (Sigma-Aldrich, St. Louis, MO, USA). We confirmed the bacterial status of each strain through PCR with forward primer 27F and reverse primer 1492R (Hoffman and Arnold 2010). We used
4 ReadyMix REDTaq polymerase for all reactions (Sigma-Aldrich, St. Louis, MO, USA). Each PCR mixture included 1 L of each primer, 12.5 L of REDTaq, 9.5 L of PCR water, and 1 L of our DNA extract, for a total of 25 L of reaction mixture per sample (Hoffman and Arnold 2010). The PCR cycling process followed Hoffman and Arnold (2010).
We used SYBR green I stain (Molecular Probes, Invitrogen) to visualize bands of DNA on 1.5% agarose gels (Hoffman and Arnold 2010). Gel electrophoresis confirmed successful amplification of bacterial DNA in all three replicates of 9133+ and 9140+. We observed no bacterial DNA amplification in replicates of 9133- and 9140-, as expected. After multiple PCRs, we were unable to successfully confirm the presence of bacteria in 9143+, so we chose not to move forward with this strain.
We cleaned the successful PCR products with 1 L of ExoSAP-IT (Affymetrix, Santa Clara, CA,
USA). The ExoSAP reaction was run in the thermocycler at 37C for one hour, and then at 80C for 15 minutes (Shaffer et al. 2017). The clean PCR products were sequenced bidirectionally at University of
Arizona Genetics Core.
We assembled the sequences of bacteria for 9133+ and 9140+ in Mesquite (Maddison and
Maddison 2009). We edited the sequences manually in Sequencher (Gene Codes Corp., Ann Arbor, MI,
USA). We compared the resulting sequences against known sequence data via BLAST (Altschul et al.
1990). We found the EHB to be Pantoea sp., as reported previously for 9140 (Araldi-Brondolo et al.
2017). However, the EHB reported previously for 9133 was a Luteibacter species, which we did not observe here. Thus, the study provided us with the opportunity to evaluate the impact of Pantoea EHB on two fungal strains.
Preparation of litter bags
We placed 3 g of autoclaved plant material of one plant species into each litter bag. Each bag was assigned to one treatment: inoculation with 9133+, 9133-, 9140+, or 9140-, or mock inoculation (control).
For each of the three plant species, we made 20 bags per treatment, for a total of 100 bags per plant species. Each bag was numbered individually.
To prepare inoculum we obtained a small plug from an actively growing culture of each fungal strain (9133+, 9133-, 9140+, 9140-) and ground it with a sterile pestle in 1 mL of sterile water in a sterile
5 Eppendorf tube. We poured each aliquot evenly over the surface of plant material in each litter bag to inoculate the material. We then added 2 mL of sterile water to the aliquot tube and poured that over the contents of each litter bag. For controls, we used only a plug of sterile 2% MEA, instead of fungal inoculum. After the bags were filled with plant material and had been inoculated, they were stapled shut, weighed individually, and wrapped in aluminum foil until deployment.
Deployment of field experiment
In December 2018 we deployed 300 litter bags into the field. We placed 50 bags of each species into each of two plots (150 bags/plot). We buried the bags at the University of Arizona Campus Agricultural
Center in two plots: one an area of native vegetation along the raised banks of a riparian area, and one in a degraded agricultural soil into which widely spaced, non-native trees were planted (Kissell, unpublished data). The plot with more natural vegetation was on a slight incline with looser, more fertile soil. The plot in the degraded soil was rocky and dry. We arranged both plots so that each would receive both sunlight and shade throughout the day. The bags were buried in shallow holes and completely covered with soil.
We ensured that each bag was ca. 10 cm away from the other bags, and no bags from the same treatment groups were next to each other in the plot. Both plots were approximately 2.5 x 2.5 m. We marked the perimeters of the plots with flags and left the bags completely buried until March 2019.
Mass-loss analysis
In March 2019 we retrieved the litter bags from the field. When the bags were completely dry, we used a toothbrush to clean soil off of each bag. The bags were individually weighed. To analyze our data, we first determined the mass loss for each bag (original weight – final weight). We then divided that value by the original weight to calculate percent mass loss (mass loss score). We scaled the mass loss scores by the average of the controls from the same plant species in the same plot. We therefore present the results as scaled mass loss for each plant species and treatment in each plot. A scaled mass loss score = 1 indicates that the percent mass loss from treatments did not differ from controls. A scaled mass loss score >1 indicates that the percent mass loss from treatments exceeded of controls, and <1 indicates that the percent mass loss from treatments was less than that of controls.
6 Results
We observed mass loss in all bags that were deployed to the field. We first evaluated mass loss from controls to determine if there were differences among plant species and plots in mass loss without our treatments. We found that control bags lost an average of 26.6% of mass (95% confidence interval: 25.2-
28.1%) over the three-month study. Mass loss values differed markedly among plant species and plots
(R-squared = 0.72, analysis of variance F (5, 51) = 29.88, P <0.0001 (Figure 1).
Figure 1. Mass loss score (percent mass loss) for control bags as a function of plot (1 = riparian, 2 = degraded) and plant species (CA, Cupressus arizonica; JD, Juniperus deppeana; PO, Platycladus orientalis).
We found that mass loss from controls varied by plant species and by plot, especially for C. arizonica. Overall, the trend was for more mass loss in the riparian plot and less in the degraded plot. Based on these differences, we used the mean values for controls for each species in each plot to produce scaled mass loss values for our treatments.
Scaled mass loss as a function of treatment
Next, we compared scaled mass loss among treatments (Figure 2). In plot 1 (riparian area) we found that all treatments were similar to controls for C. arizonica (i.e., scaled mass loss did not differ markedly from
7 1) (Figure 2). However, in plot 2 (degraded), all treatments were associated with an increase in mass loss relative to controls. There was no detectable effect of EHB in degradation of C. arizonica (Figure 2).
For J. deppeana, all treatments were associated with slight increases in mass loss relative to controls in both plots (Figure 2). There was no detectable effect of EHB in degradation of J. deppeana (Figure 2).
For P. orientalis, three treatments were associated with less mass loss than controls: 9133- (both plots)
9133+ (both plots), and 9140+ (particularly in plot 1) (Figure 2). Treatment with 9140- did not differ relative to controls. Variation in scaled mass loss reflected treatment and did not differ between plots
(multiple regression of scaled mass for P. orientalis as a function of treatment and plot (R-squared = 0.71,
F (4,72) = 44.29, P<0.0001; treatment effect, P<0.0001, plot effect, P = 0.4802). Post-hoc tests indicate more mass lost from the EHB- strains relative to the EHB+ strains of each fungus (P<0.05). Overall these results show that scaled mass loss was generally decreased by treatment in P. orientalis relative to controls, and mass loss was particularly decreased by treatment with EHB+ strains.
Figure 2: Scaled mass loss in three plant species (CA, Cupressus arizonica; JD, Juniperus deppeana; PO, Platycladus orientalis) treated with each of two species of plant-associated fungi (9133, 9140) with EHB (+) or without EHB (–). Mass loss scores ([original mass-final mass]/original mass) were scaled by dividing each mass loss score by the mean for controls of that plant species in that plot.
8 Discussion
Many plant-associated fungi form symbiotic relationships with endohyphal bacteria (EHB), which live within the healthy tissue (mycelium) of some fungi. Previous research has shown that some EHB can influence cellulase and ligninase activity of certain fungi in vitro. The goal of this project was to evaluate the degree to which EHB influence the ability of such fungi to degrade plant material under field conditions. We anticipated that mass loss would differ among plant species and as a function of fungal treatment, and that EHB would influence mass loss in a manner consistent with their previously observed results in vitro. A pilot project and in vitro trials suggested that EHB were associated with increased tissue degradation and mass loss relative to the same fungi without EHB, but our results did not meet this expectation.
Our results suggest that EHB were not associated with greater degradation of plant material. In one species an effect of EHB was detected; in that case (P. orientalis), EHB- samples lost more mass than EHB+ samples. However, we take care not to conclude that EHB- treatments were increasing tissue degradation per se: EHB- treatments either resembled controls (9140-) or had less mass loss than controls (9133-). In those cases, EHB+ treatments also decreased mass loss relative to controls. We interpret this result as suggesting that the naturally occurring organisms most capable of degrading P. orientalis were inhibited somewhat by 9140+, to a greater degree by 9133-, and especially by 9133+. This encourages further exploration as to whether 9133+ is particularly inhibitory to the subset of soil organisms that degrade P. orientalis. Interestingly, 9133+ did not have a similar effect on mass loss in the other two plant species. We predict that the community of organisms that degrades P. orientalis in these soils is somewhat unique from that degrading C. arizonica and J. deppeana, perhaps consistent with the non-native status of P. orientalis in the region. This is a topic for further study.
It is possible that the rainy, cool weather in this trial, which differed from our pilot (cool but very dry), can explain the different results relative to a pilot study that previously showed that EHB were associated with more mass loss from conifer foliage in field conditions. Tucson received an abnormally high amount of rain in winter 2018-2019 (approximately 91mm from December 2018-March 2019). The large amount of rain coupled with cool temperatures allowed an abundance of weeds to flourish, particularly in plot 1 (riparian). Rain can impact the functional activity soil microbes (Kardol et al. 2010)
9 and may have influenced our mass loss results. Specifically, the excess rain and a subsequent change in soil microbial community (Classen et al. 2015) may have diluted the effect of EHB on mass loss to an undetectable level in this study, while also allowing EHB- fungi to degrade plant material at a much higher rate than expected. In future work we suggest retrieving samples at staggered dates before 3 months when the season is particularly rainy. It is plausible also that the results we observe reflect microbial interactions, with the focal fungi (especially with EHB) possibly repelling certain fungi in soil with degradative ability. If so, then variation in such effects among plants species (see Figures) suggests that soil borne microbes that could degrade plant material differed among plant species, per above (see also discussion of plant traits, below).
Differences between plots
Overall, we observed greater mass loss for controls deployed in the riparian plot (plot 1) vs. the degraded, post-agricultural plot (plot 2). Plot 1 contained native plants and moist, relatively fertile soil. Microbial communities found in this type of soil often express more nutrient cycling genes and genes associated with breaking down plant-derived compounds (Fierer et al. 2012). In turn, communities found in arid, desert soil tend to express genes necessary for dormancy and osmoregulation, and fewer genes used in catabolism of plant-derived compounds (Fierer et al. 2012). Our controls suggest that a disparity in the composition or functional aspects of microbial communities in each plot could account for some of the difference in mass loss between plots 1 and 2. Notably, foliage of C. arizonica treated with fungi lost more mass in plot 2 than in plot 1, consistent with our prediction that the fungi and EHB considered here were interacting with soil borne microbial communities.
Differences among plant species
Plant species varied in the amount of mass loss observed (see Figure 1). This could be due in part to differences in oils and antimicrobial compounds found in each plant. Cupressus arizonica expresses high levels of monoterpene hydrocarbons in its branches and large amounts of oxygen-containing monoterpene hydrocarbons in leaves. The leaf oil of C. arizonica is rich in -pinene and umbellulone.
These oils have mild levels of antimicrobial activity against Escherichia coli, Staphylococcus aureus,
10 Enterococcus faecalis, Streptococcus pneumoniae, and Klebsiella pneumoniae (Cheraif et al. 2007).
Juniperus deppeana produces high levels of oxygen-containing monoterpenes including camphor and linalool (Adams et al. 1984). This species has hexane and methanol soluble extracts in its leaves and bark, which exhibit antimicrobial activities in vitro (Clark and McChesney 1990). The majority of the oils produced in P. orientalis are monoterpene hydrocarbons, with oxygen-containing monoterpene hydrocarbons representing a low percentage of the oils produced in the fruit and leaves of this species.
The essential oils found in the leaves of this plant species do not exhibit antimicrobial activity, while oils in
P. orientalis fruit have low levels of antimicrobial activity against Bacillus subtilis, Candida albicans, E. coli, and S. aureus. (Hassanzadeh et al. 2001). Litter bags containing P. orientalis had the highest percentage mass loss in both plots (Figure 1, Figure 2). The relative lack of antimicrobial activity in P. orientalis leaf material could account for this observation, as resident soil microbes would have been less inhibited by this plant relative to the others considered here.
Conclusions
We found that leaf material of three tree species differed in the amount of mass loss during a field experiment. The amount of mass loss also differed between plots with different soil types. In contrast to expectations, EHB were not associated with increased mass loss from plant tissue, suggesting the potential biotic interactions between EHB/their host fungi and soil microbes, especially as showcased by
P. orientalis. Future work should harvest bags at earlier intervals, evaluate soil microbial communities and their functional elements with respect to plant tissue degradation, and determine whether in vitro enhancement of cellulase and ligninase activity by EHB can be temperature- or otherwise environmentally dependent.
Acknowledgments
First and foremost, thank you to Dr. A. Elizabeth Arnold for sharing her knowledge and experience with me, as well as for being a mentor to me during my years in her lab. Throughout my time there, she has always been willing to share her wisdom, personal time, and resources to allow me to grow as a scientist and as a student. Without Dr. Arnold’s patience and dedication, this project would not have happened.
11 Thank you to Ming-Min Lee, the lab manager of the Arnold Lab. Ming’s endless patience, support, and commitment to the success of everyone in this lab has helped shape countless projects, including my thesis. Her guidance was instrumental in developing the techniques utilized in this project.
Thank you to Desirae Kissell and Ashton Leo for their assistance with the preparation of the materials used in this project and for continuing to make me feel welcome in the Arnold Lab. Additionally,
I would like to thank Joe Spraker for allowing me to use the fungal samples he had meticulously preserved and stored. Access to these samples allowed this project to move forward.
Thank you to the University of Arizona Campus Agricultural Center for allowing me to utilize the facilities and land there during my thesis.
Finally, thank you to all the members of the Arnold Lab for being kind, welcoming, and always willing to lend their assistance and expertise. Every person in this lab has made my research experience enjoyable and rewarding.
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