ABSTRACT
Biological soil crusts (biocrusts) are globally important microbial communities inhabiting the top layer of soils. They provide multiple services to dryland ecosystems but are particularly vulnerable to anthropogenic disturbance from which they naturally recover only slowly. Assisted inoculation with cyanobacteria is held as a promising approach to promote biocrust regeneration. Two different methodologies have been developed for this purpose: mass cultivation of biocrust pioneer species (such as the cyanobacteria Microcoleus spp.) on cellulose supports, and polymicrobial cultivation of biocrusts in soils within greenhouse settings. Here, we aimed to test a novel method to grow cyanobacterial biocrust inoculum based on fog irrigation of soil substrates (FISS) that can be used with either culture-based or mixed-community approaches. We found that the FISS system presents clear advantages over previous inoculum production methodologies; overall, FISS eliminates the need for specialized facilities and decreases user effort. Specifically, there were increased microbial yields and simplification of design compared to those of the culture-based and mixed-community approaches, respectively. Its testing also allows us to make recommendations on underexplored aspects of biocrust restoration: (i) field inoculation levels should be equal to or greater than the biomass found in the substrate and (ii) practices regarding evaluation of cyanobacterial biomass should, under certain circumstances, include proxies additional to chlorophyll a.
IMPORTANCE Biocrust inoculum production for use in dryland rehabilitation is a powerful tool in combating the degradation of dryland ecosystems. However, the facilities and effort required to produce high-quality inoculum are often a barrier to effective large-scale implementation by land managers. By unifying and optimizing the two foremost methods for cyanobacterial biocrust inoculum production, our work improves on the ease and cost with which biocrust restoration technology can be translated to practical widespread implementation.
INTRODUCTION
Biological soil crusts (biocrusts) are complex communities of microorganisms occurring in the top layer of arid soils, found most commonly in the plant interspaces of dryland ecosystems (1, 2). Though their biological composition can vary, biocrusts typically consist of photosynthetic primary producers, heterotrophic bacteria (3), archaea (4), and fungi (5). Most commonly, the phototrophs are cyanobacteria (6), but are sometimes also eukaryotic algae (7) and, in well-developed crusts, mosses (8) and lichens (9). The predominant cyanobacteria in arid land biocrusts, the filamentous bundle-forming Microcoleus spp. in particular, are both the main primary producers and usually the first colonizers of bare soils (10). Biocrusts provide essential ecosystem services such as soil stabilization against erosion (11) and soil fertilization contributing to carbon and nitrogen inputs (12). The initial stabilization by pioneers allows other key biocrust organisms, such as heterocystous cyanobacteria (13), lichens, and mosses, to secondarily colonize. However, biocrusts are most fragile when dry and become vulnerable to compressive forces caused by human activity, such as cattle grazing, recreational vehicles, and foot traffic (14, 15). These disturbances along with sand deposition (16, 17), prolonged drought (18), and precipitation changes brought about by climate change (19) can lead to destruction of biocrust cover, contributing to soil loss and the formation of fugitive dust, which becomes an environmental health hazard (20). Because biocrust communities are only metabolically active when hydrated (21–23), disturbed areas recover slowly, a process that can take anywhere from a few years to decades or centuries (24, 25).
Therefore, efforts to speed biocrust recovery have recently become an important focus in dryland ecological restoration. Initial studies transplanted healthy biocrusts to disturbed areas, demonstrating that restoration is possible (24, 26, 27). However, attempts to restore disturbed areas at large scales by using healthy crust is not sustainable and does not guarantee success (28). Current methods of biocrust restoration have shifted the focus to artificially producing keystone phototrophic biocrust components (cyanobacteria, lichens, or mosses) for assisted inoculation. Many proposed methods to scale up biocrust components, such as lichens and mosses, show promise (29–31). However, the scale-up of cyanobacterial components, such as pioneer Microcoleus spp., continues to bottleneck large scale restoration efforts. The key cyanobacterial pioneers of biocrusts, Microcoleus vaginatus and Microcoleus steenstrupii, are difficult to grow using traditional liquid-based cyanobacterial scale-up methods (32) and are prone to contamination by weedy species in open systems (33). Two studies have recently advanced biocrust inoculum production: the “cellulose tissue support” (32) and the “mixed-community nursery-based” methods (34). The cellulose tissue support method involves scale-up of filamentous cyanobacterial biomass isolated from remnant biocrust on floating cellulose tissue under sterile conditions. While inherently successful in producing site-specific genetically controlled inoculum, it is labor intensive and relies on specialized equipment, facilities, and personnel. The cellulose tissue support method also produces inoculum that is not acclimated to desiccation or high light intensities, requiring implementation of “hardening” protocols to increase inoculum performance in the field (35), further adding to time and labor costs. The mixed-community nursery-based approach (34) includes an initial fractional factorial analysis to optimize conditions for rapidly producing large quantities of biocrust inoculum from biocrust remnants on native soils without allowing the overall native cyanobacterial community composition to shift significantly. While this method can produce high yields, it requires site-specific experimentation, is subject to ravages by biocrust pathogens (36), and may result in inoculum adapted to optimal growth conditions rather than to comparatively harsher field conditions. It also requires a complex wicking-based watering system (37), constant monitoring, and fine-tuning (34, 36), restricting its potential use in remote locations. Further optimization of these methods is needed to make large-scale restoration efforts more approachable for land managers. Here, we present a modification of the methods described above to unify and optimize the production of cyanobacterial biocrust inoculum. We developed and tested a flexible fog-based irrigation system (FISS) to successfully scale up both cultured cyanobacterial pioneer organisms and polymicrobial biocrust communities on native soils. The FISS method unifies biocrust inoculum production methods, increases yields and fitness to field-like conditions, and simplifies production by eliminating the need for special infrastructure.
RESULTS
FISS growth of cultures.We tested the integration of culture-based inoculum production with the FISS system (Fig. 1) on native soils as the substrates, under both laboratory (indoor) and local outdoor conditions. Under indoor conditions, M. vaginatus HSN003 and FB020 yielded 43.57 and 22.17 mg chlorophyll (Chl) a m−2, respectively (Fig. 2), and M. steenstrupii HS024 and JS010 yielded 43.36 and 14.9 mg Chl a m−2, respectively (Fig. 2). Biomass yields under outdoor conditions were overall greater than those indoors: M. vaginatus HSN003 and FB020 yielded 94.56 and 30.06 mg Chl a m−2, respectively (Fig. 2), and M. steenstrupii HS024 and JS010 yielded 84.80 and 23.34 mg Chl a m−2, respectively (Fig. 2). Differences between treatments were significant for the cold desert strains (t test, HS024, P < 0.001; HSN003, P < 0.005) but nonsignificant for the hot desert strains (t test, JS010, P = 0.17; FB020, P = 0.09). Full data sets of Chl a concentrations and visual aspects can be found in Table S1 and Fig. S1, respectively, in the supplemental material. Microscopic examination of culture plate and control plates showed no cyanobacterial contaminants in indoor plates across all time points. The presence of noninoculated cyanobacteria was detected at low abundance in the last time point of the outdoor sterilized control plates.
(A) Experimental set up of FISS chambers during the outdoor growth trials on the roof of the Life Sciences Building East at Arizona State University (Tempe, AZ) during October to November 2018. (B) Automated scale-up of FISS chamber to 1.5-m plastic pools with cotton fitted sheet covering. (C) General schematic of FISS system setup.
Growth dynamics (mean biomass ± standard deviation [SD], n = 12) of FISS-grown Microcoleus cultures. Shaded lines indicate maximum average Chl a values (±SD) obtained with the floating cellulose tissue method (32) for the same strains at peak biomass.
FISS growth of mixed communities.We substituted the wicking-based watering system used previously in the mixed-community nursery-based approach (37) with the FISS system. The final average Chl a content after 14 wetting events was significantly higher than inoculation contents (Fig. 3) (analysis of variance [ANOVA], hot desert, P < 0.001. cold desert, P < 0.001) in all cases (shaded and nonshaded treatments and hot and cold desert original locations). While average final Chl a content tended to be higher in the shaded than in nonshaded treatments (Fig. 3), the differences were not significant in either soil (ANOVA, hot desert, P = 0.79; cold desert, P = 0.11). In both locations, control plates also showed increases in final average Chl a over time, suggesting that growth of noninoculum photosynthetic organisms already present in (unsterilized) soil substrates had taken place (Fig. 3). This was significant in the cold desert (t test, P = 0.03) but not in the hot desert soils (t test, P = 0.09).
(A) Visual aspect of mixed-community, outdoor-incubated FISS-grown biocrusts under different shade treatments. (B) Chlorophyll a concentrations in mixed-community FISS trials given in mg Chl a m−2. Shown are means (n = 5) ± standard deviations. P values are the result of an ANOVA between treatment Chl a values (95% confidence interval).
To confirm Chl a-based biomass determinations, and because we expected variability in cellular Chl a content in response to differing light intensities used, quantitative real-time PCR (qPCR) determinations of 16S rRNA copy numbers were carried out (see Table S2). As expected, 16S rRNA-based values for absolute cyanobacterial abundance did not always correlate to Chl a content (Fig. 3). In agreement with Chl a data (Fig. 3), FISS-grown biocrusts from the cold desert had, on average, fewer 16S rRNA gene copies than field biocrusts (Table S2). However, in contrast with Chl a data, the nonshaded treatments had significantly more 16S rRNA gene copies than the shaded treatment (ANOVA, P = 0.046), similar to those found in the field (ANOVA, P = 0.14), suggesting that cyanobacteria in nonshaded treatments were less pigmented. Here, the final 16S-based biomass was equivalent to that of established field biocrusts. In hot deserts, 16S-based biomass yield from both nonshaded and shaded treatments (ANOVA, P = 0.02 for both) were significantly higher (approximately 4-fold) than those in original field biocrusts. Shaded and nonshaded treatments from the hot desert contained similar numbers of 16S rRNA gene copies (ANOVA, P = 0.36).
Mixed-community bacterial composition.The bacterial community compositions of the original remnant biocrusts (field) were typical for their respective locations (13, 19, 38), with Cyanobacteria, Proteobacteria, Bacteroidetes, and Actinobacteria being the major components of the community in both locations (see Fig. S2 and S3). In cold desert biocrust, a majority of the cyanobacteria consisted of Microcoleus vaginatus, members of the M. steenstrupii complex, and Tolypothrix spp., with M. vaginatus and M. steenstrupii complex together accounting for more than 70% of total cyanobacterial reads (Fig. 4B and S4). In hot desert biocrusts, a majority of the cyanobacterial composition consisted of M. vaginatus, M. steenstrupii complex, and Scytonema spp., with M. vaginatus and M. steenstrupii complex together accounting for more than 50% of total cyanobacterial reads (Fig. 5B and S5). In these locations, M. vaginatus was dominant in cold desert locations and M. steenstrupii complex was dominant in hot desert locations, as is consistent with previous reports on geographical dominance partitioning (39).
(A) NMDS ordination of Bray-Curtis pairwise distance computed on the cyanobacterial sequence composition of remnant cold desert biocrust samples (field), native soil (substrate), or FISS-grown mixed-community inoculum (shaded/nonshaded), with 95% confidence ellipses drawn for each with a stress value of 0.049. Cyanobacterial communities show significant differences between treatments (PERMANOVA, P < 0.001). (B) Cyanobacterial abundance and community structure of cold desert biocrusts of FISS-grown inoculum under different shade treatments, as determined by high-throughput 16S rRNA gene analysis coupled to qPCR. Note: “Microcoleus steenstrupii” corresponds phylogenetically to a complex of several genus-level clades (19), these clades are distinguished in the figure.
(A) NMDS ordination of Bray-Curtis pairwise distance computed on the cyanobacterial sequence composition of remnant hot desert biocrust samples (field), native soil (substrate), or FISS-grown mixed-community inoculum (shaded/nonshaded), with 95% confidence ellipses drawn for each with a stress value of 0.049. Cyanobacterial communities show significant differences between treatments (PERMANOVA, P < 0.001). (B) Cyanobacterial abundance and community structure of hot desert biocrusts of FISS-grown inoculum under different shade treatments, as determined by high-throughput 16S rRNA gene analysis coupled to qPCR. Note: “Microcoleus steenstrupii” corresponds phylogenetically to a complex of several genus-level clades (19); these clades are distinguished in the figure.
The cyanobacterial community composition of cold desert FISS biocrusts, in both shaded and nonshaded treatments, was significantly different from that of their original community (Fig. 4A) (permutational multivariate analysis of variance [PERMANOVA], P = 0.001 and 0.002, respectively). Overall, in the cold desert, the nonshaded treatment matched the field cyanobacterial community better in terms of presence and absolute abundance of Microcoleus, Tolypothrix, and Scytonema (Fig. 4B and S4). In hot desert FISS biocrusts, cyanobacterial community compositions of shaded and nonshaded treatments were significantly different from those of the field (PERMANOVA, P = 0.001 for both), with the shaded treatment closely resembling the community composition found in the noncrusted soil substrate (PERMANOVA, P = 0.10) (Fig. 5A). The cyanobacterial genera Potamolinea, Leptolyngbya, and Nostoc were major components of the cyanobacterial community found in the noncrust substrate and in the shaded and nonshaded FISS biocrusts but absent or rare from the field biocrust community (Fig. 5B and S5).
Overall, nonshaded treatments differentially promoted the growth of many cyanobacterial taxa relevant to the original biocrust community in both locations, based on fold change of 16S rRNA gene copy numbers. In cold desert FISS biocrusts, all relevant cyanobacterial taxa had greater growth under nonshaded than under shaded conditions (see Fig. S6). To a lesser extent, in hot desert FISS biocrust, nonshaded treatments promoted the growth of relevant cyanobacterial taxa such as Microcoleus spp. (see Fig. S7).
DISCUSSION
Culture- and mixed-community FISS scale-up.We aimed to improve upon established methods for biocrust inoculum production by using an ultrasonic mist maker that fills a semi-enclosed transparent chamber with fog in order to saturate native soil substrates without overwatering. We succeeded in improving upon the floating cellulose tissue method (32) that, from very little source material, was able to produce large quantities of pedigreed cyanobacterial biomass for use as restoration inoculum. Using FISS, biomass yields exceeded those produced by the floating cellulose tissue method in every case (Fig. 2). This is likely because native soil substrate has a higher capacity to support cyanobacterial biomass within their matrix than a heavily two-dimensional cellulose tissue. Additionally, cultures grown on native soils seemed to reach a carrying capacity and remain stable (Fig. 2; see also Fig. S1 in the supplemental material), unlike cellulose-grown biomass, which typically crashes shortly after achieving peak biomass (32). Furthermore, because the FISS approach works well under outdoor conditions, it renders unnecessary any “preconditioning” treatments before final inoculation, such as those previously described (32, 35) to improve field survival, thus significantly minimizing effort. However, because FISS does not use fully sterile settings, we suggest regular monitoring by microscopy to rule out contamination (as we detected in the control plates at the final time point). Though the FISS method may require more time to reach peak biomass, the cost is offset by eliminating the effort of high-volume inoculation and monitoring of cellulose filters with fickle growth dynamics. The FISS approach does away with a constant need of sterile equipment/environments, is highly portable, and can be fully automated (Fig. 1). Overall, we see this approach as a much-needed improvement.
In terms of yields, FISS also succeeded in improving upon the mixed-community nursery-based method (34) that produced high yields of inoculum compositionally similar to the native field biocrust using remnant biocrust as source material, eliminating the need for greenhouse facilities and shading, the two factors that make this approach unfeasible for widespread use by land managers. Because the FISS system is portable, easily scalable, and automatable, mixed-community scale-up can principally be carried out in climatically appropriate locales, reducing the need for constant monitoring and doing away with the need for extensive infrastructure (Fig. S1B). Furthermore, we showed that with FISS, the process can be simplified by removing cumbersome expensive shading elements, as total cyanobacterial yield, as determined by total 16S gene copy number, was, at worst, independent of this treatment (Fig. 4 and 5; Table S2). In fact, absence of shading seemed to select for motile biocrust pioneer species (Fig. S6 and S7), a desirable element in an inoculum. While the cyanobacterial community composition indeed shifted in both treatments from that of field biocrusts in the cold desert trial (Fig. 4A), there was an overall increase in the abundance of relevant cyanobacterial taxa found at that location, such as Microcoleus, Tolypothrix, and Scytonema (Fig. S6). While the issue of community stability remains an area of potential improvement, the shifts for major cyanobacterial players are minimal enough as to be inconsequential here, the inoculum maintaining the dominance of desired species, especially in the nonshaded communities.
Thus, we were successful in transferring both culture-based and mixed-community approaches to the FISS system platform, thus unifying biocrust inoculum production into a methodology that is simplified, scalable, and ready for widespread implementation. Using commercially available ultrasonic mist makers, the FISS platform can be easily deployed to a 5- to 6-m2 growth chamber. Based on our culture-based yields, 10 such FISS chambers, easily managed by a single operator, can produce quality inoculum to restore 1,000 to 2,000 m2 at 5 mg Chl a m−2 within 6 months. Such half-acre scale already has potential for applications in settings such as the restoration of semiurban developments in arid lands or small solar farms. As with other restoration methodologies, such as reforestation, it is the process of continual stepwise optimization and expansion of facilities that enables effective restoration at truly large scales.
Influences of preexisting soil substrate community and inoculation levels on FISS inoculum composition.While not an intended part of our experimental design, we observed that the cyanobacterial community composition of hot desert biocrusts, for both shaded and nonshaded treatments, differed significantly from that of the original field biocrust (Fig. 5A). Surprisingly, the typical pioneer cyanobacterium, Microcoleus, was almost entirely absent from both shaded and nonshaded communities despite being the original dominant genus in field communities (Fig. 5B and S5). Additionally, cyanobacterial taxa not typically dominant in biocrust communities from this location, such as Potamolinea, Leptolyngbya, Oscillatoria, Pseudanabaena, and a cyanobacterium from the family Nostocaceae, appeared in abundance in our treatments (Fig. 5B). This led us to examine the cyanobacterial community that was originally present in the bulk soil we used as the substrate. Indeed, particularly in these hot desert sites, there was no significant difference between soil substrate community and shaded treatment communities (PERMANOVA, P = 0.10) (Fig. 5B). This suggests that soil substrate community composition should be examined and taken into consideration when evaluating population growth dynamics and when determining the level at which to inoculate soil substrates used in restoration. For both hot and cold desert locations, we inoculated at 5% of the original, which is equivalent to inoculation concentrations of 1.1 and 5.6 mg Chl a m−2, respectively. Baseline Chl a concentration in the soils used as growth substrates was 3.5 mg Chl a m−2 for the hot desert and 5.6 mg Chl a m−2 for the cold desert. Thus, in the hot desert FISS, inoculum was only approximately one-third of the existing cyanobacteria, whereas in the cold desert FISS, it was equal (Fig. 3). Clearly, this may have played a role in the compositional outcomes, particularly when inoculation level was relatively low. This suggests there may be a minimum level of biomass inoculation required to overcome the growth potential of bacterial communities found in the soil substrate if the soils are not sterilized prior to seeding. When using nonsterile soil substrates for biocrust inoculum cultivation, it would be desirable to inoculate at levels higher than those found in soil substrates.
Alternatively, of course, one could sterilize the soil substrate, but we see this as a significant burden to scalability.
Limitations of Chl a as a proxy for photosynthetic biomass.Chl a is commonly used to determine photosynthetic biomass of oxygenic phototrophs, despite also being described as a poor proxy (40). The reasoning is that phototrophic organisms are able to modulate their Chl a content in response to changes in growth conditions, chiefly, light intensity (41). For example, phototrophic organisms grown at low light (shade types) intensities contain more Chl a per cell to capture limiting light than the cells of the same organism grown at high light (sun types). In comparing Chl a-based biomass, one should be aware of this potential interference. Our experiments, which involved variations in light exposure levels, provide additional evidence for this effect, so that in shade treatments, Chl a was a significant overestimator of biomass at both locations relative to those of 16S-based proxies. Similarly, relying on visual observations of “biocrust cover” by color may not accurately reflect phototrophic biomass, because in addition to the variations in pigment content, motile cyanobacteria, such as the pioneer Microcoleus spp., move in the soil profile to optimal light irradiance (42). At high light intensities, these cyanobacteria move down into the soil, becoming less visible, to the point that the soil may appear bare, as seen in Fig. 3. We recommend that a biomass estimator other than visual observation or Chl a be used in experiments involving changes in light intensity.
Conclusion.The FISS approach for biocrust inoculum production represents significant improvement modifications of two current methods, the cellulose tissue support (32) and the mixed-community nursery-based (34) methods, increasing yields while significantly reducing labor costs in scale-up and conditioning, eliminating the need for infrastructure, and adding portability. Our experiments also provide evidence that inoculation level and soil substrate communities have an influence on the final community composition of biocrust inoculum that should be taken into account and that Chl a can be an unreliable proxy for phototrophic biomass in biocrusts when changes in light intensity are involved in experimentation.
MATERIALS AND METHODS
Biocrust and isolate sourcing.Experiments were performed with biocrust remnants and cyanobacterial isolates from four U.S. southwest locations with different climatic and edaphic characteristics; hot desert biocrusts and soil substrates were collected from the Chihuahuan Desert (south Texas, Fort Bliss military base; lat 32.431069°, long −105.984151°), and cold desert biocrusts and soil substrates were collected from the Great Basin Desert (Hill Air Force Base-Utah Test and Training Range, lat 41.104198°, long −113.008204°). Additional location and sample details are described in previous work (34). Cyanobacterial isolates used were originally isolated from remnant biocrust in those locations: four pedigreed Microcoleus species isolates (M. vaginatus FB020 and M. steenstrupii JS010 isolates from the hot desert and M. vaginatus HSN003 and M. steenstrupii HS024 from the cold desert). Additional cyanobacterial isolate details are given in previous work (32).
Culture-based FISS scale-up.The chambers used consisted of two clear plastic storage containers (Sterilite, 90 cm by 42 cm by 15 cm) arranged to form a chamber. It was provided with drainage holes at the bottom and an open window on top (Fig. 1). A smaller plastic container (Sterilite, 36 cm by 20 cm by 12 cm) placed inside served as a water reservoir. A commercial ultrasonic mist maker (5-disc; The House of Hydro, Ft. Myers, FL) was placed in the reservoir. All materials were sterilized with 70% ethanol and 1 h of UV exposure prior to set up, and the reservoir was filled with reverse osmosis pure water (RO water). Native soil from each location was autoclaved five times (30 min, 121°C), and aliquoted into 8 sterile (1-cm by 10-cm diameter) petri plates per soil type, saturated with Jaworski’s minimal medium (JM) (43). Due to the clumping tendency of Microcoleus strains (32), each strain used was homogenized by forcing the culture through a sterile 60-ml syringe until observed clumps were broken apart (32, 35). Each homogenized strain was then inoculated separately on JM saturated soil substrate to a final concentration of 0.8 to 5 mg Chl a m−2 depending on the initial concentration of a given strain. Four plates of each isolate were then placed uncovered into outdoor and indoor FISS chambers and allowed to air dry for 48 h. The outdoor fog chamber was covered with a shade cloth that blocked 60% of incoming solar radiation. The indoor chamber was under 14 h of illumination (19 to 21 microeinsteins m−2 s−1; Philips fluorescent lamp) and 10 h of darkness daily at an average indoor temperature of 23°C. In outdoor trials, mean temperatures ranged from 17 to 23°C. Plates were then wetted during incubation by turning on the ultrasonic mist maker for 15 min, at which time, the soil substrate was saturated and a thin water film was observed on the surface. After 24 h of wet conditions, plates were allowed to dry and remained dry for 48 h. This watering cycle was repeated 12 times, refilling the reservoir with RO water as needed. Uninoculated JM-saturated plates of each soil were used as negative controls. Plates were sampled for growth initially, 24 h after wetting, and after 3, 6, 9, and 12 wetting events. Chl a sampling was performed by randomly collecting three (1-cm diameter, 0.5-cm deep) cores from each plate, which were stored at 4°C in darkness until extraction. At each sampling time, random microscopic inspections of control and inoculated plates were performed to ensure growth was not due to aeolian contamination.
Mixed-community FISS scale-up.Here, unsterilized native soil substrate was aliquoted into sterile petri plates (1-cm by 15-cm diameter; VWR). Remnant biocrusts from each location were lightly homogenized by disruption with a mortar and pestle and inoculated onto ten replicate native soil substrate plates at 5% concentration of the original remnant biocrust (based on areal Chl a concentration). Outdoor growth trials for both hot and cold desert biocrusts were performed in the fall, avoiding summer heat, as previously suggested (35, 44). Mean temperatures ranged from 17 to 23°C and light intensity peaked at ∼1,800 microeinsteins m−2 s−1. Biocrust-inoculated plates from both hot and cold desert locations were divided into two FISS chambers and subjected to a wetting event every 3 days for 14 wetting events. Each plate from both locations received 1 ml of a phosphorus solution (80.5 mM K2HPO4, 80.6 mM KH2PO4), and additionally, hot desert plates also received 1 ml of a nitrogen solution (468 mM NH4NO3) during the initial watering event. One growth chamber was covered with 60% shade cloth based on the optimization for each location as previously described (34) (n = 5), and in a second FISS chamber, unshaded growth trials were also performed for each location to ascertain the effect of increased light and UV exposure on biomass yield and community composition (n = 5). Control plates (n = 3 for each location) consisted of uninoculated unsterilized soil substrate with phosphorus and nitrogen additions as described above. For both initial and final time points, 20 (1.2-cm diameter, 1-cm deep) cores per plate were collected, dried, and pooled into a single sample in 50-ml centrifuge tubes (Falcon) and stored at −80°C until Chl a and DNA extractions were performed. Additionally, 10 (1.2-cm diameter, 1-cm deep) cores of each unsterilized soil substrate were collected, dried, and pooled in 50-ml centrifuge tubes (Falcon) and stored at −80°C for eventual Chl a and DNA extractions.
Microbial community composition.Pooled cores from each mixed-community plate (20 cores per plate) were lightly homogenized with mortar and pestle. A weighted aliquot of this pooled sample was used for DNA extraction. The same process was repeated with the uninoculated controls. Soil DNA was extracted with a PowerSoil extraction kit (Qiagen) using the standard protocol. Bacterial/archaeal community analysis was performed via commercial next-generation sequencing in a MiSeq Illumina platform. Amplicon sequencing of the V4 region of the 16S rRNA gene was performed with barcoded primer set 515F/806R (45) according to the Earth Microbiome Project (EMP) protocol (46) for library preparation. PCR amplifications were conducted in triplicates and then pooled and quantified using a Quant-iT PicoGreen dsDNA assay kit (Invitrogen). Two hundred forty nanograms of DNA of each replicate was pooled and cleaned using a QIAquick PCR purification kit (Qiagen). The pooled DNA was quantified using an Illumina library quantification kit, ABI Prism (Kapa Biosystems), diluted with NaOH to a final concentration of 4 nM, denatured, and diluted to a final concentration of 4 pM; then, 30% of PhiX was added to the solution. The library was then loaded in the sequencer using chemistry version 2 (2 × 250 paired-end) and according to the manufacturer’s specifications (46). Sequencing was performed in the Microbiome Analysis Laboratory at Arizona State University (Tempe, AZ, USA), yielding raw FASTQ sequence files.
Bioinformatics analysis.The raw FASTQ file was demultiplexed within the MiSeq Illumina workflow under default parameters. Paired sequences were demultiplexed and analyzed via Qiime2.10 (47), using the DADA2 plugin (48) where sequences were trimmed to include 250 bases from the V4 region, bound by 515F/806R primers (49), to create a feature table with representative sequences (features) and their frequency of occurrence. To remove highly variable positions, sequences were aligned with the MAFFT program (50). FastTree (51) was used to generate a tree. Taxonomy was assigned with the naive Bayes classifier trained on the Greengenes 13.8 release. While all phyla were analyzed, additional steps were taken to identify cyanobacteria due to the poor taxonomic resolution obtained with Greengenes (52). Cyanobacterial sequences were filtered from the feature table, and cyanobacterial sequences that attained at least 0.05% of the total number of cyanobacterial features were then phylogenetically assigned to the level of greatest taxonomic resolution, generally, genus level, using our own curated cyanobacteria database/tree version-0.22a (https://github.com/FGPLab/cydrasil/tree/0.22a) via RAxML (53) and displayed using ITOL (54). For microbial community analyses, significance in composition shifts was tested with PERMANOVA calculated on Bray-Curtis similarity matrices of relative abundances derived from sequencing with 9,999 permutations and visualized using two-dimensional (2-D) nonmetric multidimensional scaling (NMDS) plots. All calculations were performed using R (55) except for shifts in specific cyanobacterial community members. Shifts in cyanobacterial community members were analyzed with SIMPER analyses using relative abundance at the genus level within the PRIMER software, v6 (56). For analyses on comparisons of Chl a content and total bacterial abundance, outliers were removed, and statistical tests were performed using R (55).
Chl a determinations.Chl a was used as a proxy for photosynthetic biomass. Three pooled cores were extracted per plate by grinding the soil-cyanobacteria mixture in 90% acetone with mortar and pestle for 3 min, as previously described (32, 35, 44). Extracts were then transferred to a 15-ml Falcon tube where the volume was adjusted to 10 ml with 90% acetone, vortexed for 30 s, and then stored in the dark for 24 h at 4°C. Absorbance spectra were recorded on a UV-visible spectrophotometer (Shimadzu UV-1601). Interference from scytonemin and carotenoids was corrected for using a trichromatic equation (57). For the cyanobacterial isolates, independent determinations were performed in triplicates per plate at each sampling time point. Chl a yield was calculated by subtracting initial values from values at the time point of interest.
16S rRNA gene copy number determinations.Absolute abundance of bacteria, used as an additional proxy for biomass, was determined by qPCR (quantitative real-time PCR) using aliquoted DNA extracted from homogenized cores for each treatment. After fluorometric determination of DNA concentration in the extract (Qubit; Life Technologies, NY, USA), we used qPCR with a universal (bacterial/archaeal) 16S rRNA gene primer set (338F, 5′-ACTCCTACGGGAGGCAGCAG-3; 518R, 5′-GTATTACCGCGGCTGCTGG-3′), which produces amplicons of ideal size for qPCR, to determine the number of 16S rRNA gene copies present in each extract. The PCR was performed in triplicates using the Sso Fast mix (Bio-Rad, Hercules, CA, USA) under conditions previously published (58). The final 16S rRNA gene copy number per unit volume of biocrust was determined from the qPCR data (copies/extract) and the total soil volume used for extraction. The number of 16S rRNA genes obtained by qPCR was later used to arrive at total population sizes for each phylum or cyanobacterial taxon by multiplying the total number of genes by the relative abundance of the taxa, as determined by Illumina sequencing and bioinformatic analyses as previously described (19).
Data availability.Raw sequence data have been submitted to NCBI and are publicly available under BioProject number PRJNA600270.
ACKNOWLEDGMENTS
We thank Julie Bethany and Phillip Heinen for assistance in experimental set up and monitoring.
C.N., A.G.-S., and F.G.-P. conceived the research; C.N. and A.G.-S. designed and performed experiments; C.N. analyzed data; C.N., A.G.-S., and F.G.-P. discussed results; and C.N. and F.G.-P. wrote and edited the manuscript.
We declare no conflict of interest.
FOOTNOTES
- Received 12 March 2020.
- Accepted 23 April 2020.
- Accepted manuscript posted online 1 May 2020.
Supplemental material is available online only.
- Copyright © 2020 American Society for Microbiology.
