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Desiccation of ecosystem-critical microbialites in the shrinking Great Salt Lake, Utah (USA) [1]

['Carie Frantz', 'Department Of Earth', 'Environmental Sciences', 'Weber State University', 'Ogden', 'Ut', 'United States Of America', 'Environmental Science Program', 'Cecilia Gibby', 'Rebekah Nilson']

Date: 2023-10

Great Salt Lake hosts an ecosystem that is critical to migratory birds and international aquaculture, yet it is currently threatened by falling lake elevation and high lakewater salinity resulting from water diversions in the upstream watershed and the enduring megadrought in the western United States. Microbialite reefs underpin the ecosystem, hosting a surface microbial community that is estimated to contribute 30% of the lake’s primary productivity. We monitored exposure, desiccation, and bleaching over time in an area of microbialite reef. During this period, lake elevation fell by 1.8 m, and salinity increased from 11.0% to 19.5% in open-water portions of the outer reef, reaching halite saturation in hydrologically closed regions. When exposed, microbialite bleaching was rapid. Bleached microbialites are not necessarily dead, however, with communities and chlorophyll persisting beneath microbialite surfaces for several months of exposure and desiccation. However, superficial losses in the mat community resulted in enhanced microbialite weathering. In microbialite recovery experiments with bleached microbialite pieces, partial community recovery was rapid at salinities ≤ 17%. 16S and 18S rRNA gene sequencing indicated that recovery was driven by initial seeding from lakewater. At higher salinity levels, eventual accumulation of chlorophyll may reflect accumulation and preservation of lake material in halite crusts vs. true recovery. Our results indicate that increased water input should be prioritized in order to return the lake to an elevation that submerges microbialite reefs and lowers salinity levels. Without quick action to reverse diversions in the watershed, loss of pelagic microbial community members due to sustained high salinity could prevent the recovery of the ecosystem-critical microbialite surface communities in Great Salt Lake.

Funding: This work was supported by National Science Foundation ( nsf.gov ) EAR # 1826869 and EAR # 1801760 to CF. The latter grant also funded the Geoscience Education Targeting Underrepresented Populations (GETUP) Summer Research Experience program at Weber State University, which supported CG, RN, MN, CE, CS, and other students mentioned in the acknowledgements. CG & RN received additional support for their work on this project from the Weber State University Office of Undergraduate Research and Department of Earth and Environmental Sciences. AS was supported by Utah NASA Space Grant #80NSSC20M0103 and a Ronald E. McNair Scholars grant #P217A220138 to Westminster College. BB was supported in part from a grant from the State of Utah, Department of Natural Resources, Division of Forestry, Fire, and State Lands. The funders had no role in study design, data collection and analysis, decision to publish, or preparation of the manuscript.

In autumn 2022, roughly 40% of the lake’s microbialites were subaerially exposed ( Fig 1D , [ 13 , 37 ]) and desiccated (Figs 1G and 3 ), representing a substantial loss of productivity. A critical question for the management of Great Salt Lake and its associated watershed is how effectively and under what conditions the microbialite periphyton communities persist. If policies are enacted that allow lake levels to rebound, do microbialite periphyton communities recover their ecosystem function? We investigated these questions during two successive summers where historic lake lowstands were reached and exceeded (2021 and 2022); this paper presents the findings along with other recent data on Great Salt Lake’s microbialite periphyton communities.

Although typically viewed as relics—geobiologic curiosities that provide a window into Earth’s deep past and life’s evolution—microbialites are beginning to gain appreciation for their role in supporting modern ecosystems (e.g., [ 25 , 26 ]). Perhaps the best-studied example of this is Great Salt Lake, where it has become apparent that the lake’s microbialites serve a critical function in the lake’s overall ecosystem. Robust photosynthetic microbial mats (periphyton, adopting for this paper the broad definition that acknowledges the presence of a diverse community, cf. [ 27 ]) exist on the surfaces of microbialites in Great Salt Lake when conditions—primarily submergence and salinity—are favorable. The periphyton is dominated by a single genus of halophilic, coccoidal cyanobacteria, Euhalothece ( Fig 2 ), although other phototrophs, including diatoms, green algae, and flagellates, can be seen associated with the mats, and the mat community is taxonomically and functionally diverse [ 12 , 28 , 29 ]. Microbialite periphyton conservatively contribute one third of the primary production in Great Salt Lake [ 30 , 31 ].They are the primary food source for brine fly (Ephydra spp.) larvae [ 32 , 33 ], as well as a seasonally important food source for the lake’s economically important brine shrimp (Artemia franciscana) [ 7 , 33 ]. Additionally, microbialites, which are presumably built by carbonate production facilitated by their periphyton [ 28 ], offer stable oases in the shifting oolitic sands and carbonate mud that compose most of the Great Salt Lake benthos. This makes them crucial habitat for Ephydra larvae, which depend on the microbialites for both food and pupation habitat [ 32 , 33 ]. The organisms that microbialite periphyton support feed in turn feed millions of birds that depend on the lake ecosystem [ 5 , 30 , 32 – 36 ] ( Fig 2A ). Lake level fall is subjecting microbialites and their periphyton to desiccation.

Low lake levels and consequent shoreline shift has exposed hundreds of square kilometers of microbialite mounds, which occur in extensive reefs [ 13 ] in Great Salt Lake’s near-shore environments and benthos. Microbialites, carbonate mounds formed by interactions of microbes with the lake’s chemical environment, are of academic interest as analogues for economically important hydrocarbon reservoirs [ 11 ] and paleoenvironmental records [ 12 , 20 , 21 ]. Microbialites are not unique to Great Salt Lake; indeed, other lakes threatened by modern lake level decline are home to similar structures, including microbially-influenced carbonate mounds in several other lakes in the Western flyway, Australia’s Lake Clifton in Australia, Ethiopia and Djibouti’s Lake Abhe, and the giant microbialites of Turkey’s Lake Van [ 22 – 24 ].

Great Salt Lake is currently threatened by a rapid decline in lake levels and consequent increase in salinity. Layered onto normal decadal cycles in precipitation [ 15 ], its watershed has been impacted by the megadrought that has gripped the western United States since 2000, which has been worsened by anthropogenic climate change [ 16 ]. An even greater threat to the lake, however, has been the overuse and diversion of the waters that would otherwise feed Great Salt Lake for agricultural, industrial, and municipal uses. Such consumptive water diversions are estimated to have reduced the lake’s volume by >60% [ 17 ]. As a result, the lake has shrunk to historic low levels in the past decade ( Fig 1E and 1F ), following a pattern of water overuse leading to lake demise seen in ecologically-important saline lakes around the world [ 1 , 5 ]. Water overuse in Great Salt Lake’s watershed has substantially impaired the lake’s resilience to future changes in regional hydroclimate [ 18 ]. It has become clear that, without an overhaul of water use policy and practice in the watershed, the lake could soon be lost [ 19 ].

(A) Simplified ecosystem diagram of Great Salt Lake’s south arm, illustrating the importance of the lake’s microbialites and the effect of microbialite exposure. Dashed arrows represent life cycle stages, solid arrows represent consumption. After Baxter (2018) [ 63 ] & Belovsky et al. (2011) [ 35 ]. (B) Stereo-photomicrograph of a Great Salt Lake microbialite piece imaged at 10x showing a healthy periphyton community, with three-dimensional clumps of Euhalothece bound by extracellular polymers, and white points of carbonate highlighted with arrows. Sample collected from Site B on July 7, 2020. (C) Phase contrast photomicrograph of a healthy microbialite periphyton community sample imaged at 400x magnification. The greenish mass is a clump of Euhalothece. Also visible in association with the Euhalothece mat are a pennate diatom (arrow 1), filamentous organism (arrow 2), and green alga (arrow 3). Sample was collected October 10, 2019 from Site B. (D) Positive phase and differential interference contrast photomicrograph imaged at 1000x magnification of Euhalothece culture from a Great Salt Lake microbialite sample collected in 2019 at Antelope Island State Park [ 14 ] .

Great Salt Lake comprises not only the hypersaline open water but also distinct habitats along a salinity gradient, including fresh- to brackish-water estuaries and wetlands where rivers enter the lake, and expansive mudflats and playas. The main body of the lake is segmented by a rail causeway, which isolates the salt-saturated north arm (Gunnison Bay) from the south arm (Gilbert Bay), which encompasses our study site. The south arm supports a relatively simple but significant food web ( Fig 2A ); Great Salt Lake is a hemispherically important ecosystem [ 4 ] that supports millions of resident and migratory birds [ 5 , 6 ] and a brine shrimp industry that harvests cysts used as feed in global aquaculture [ 7 ]. The lake is a hypersaline Cl-Na-SO 4 -Mg-dominated system [ 8 , 9 ] with chemical and biological factors contributing to its “carbonate factory” [ 10 ] despite moderate modern pH values; carbonate deposits include oolitic sand, organic-rich carbonate mud, and mounded reef-forming microbialites [ 11 , 12 ].

(A) Map of Great Salt Lake showing the north (N) and south (S) arms of the lake with major sites described in this paper: USGS lake elevation sites 1001000 (ES1, Saltair site) and 10010024 (ES2, Causeway site), weather station sites KUTSYRAC22 (WS1) and KUTSYRAC27 (WS2), Buffalo Point microbialite reef sites (BP), and Ladyfinger Point (LFP). Left inset shows the location of Great Salt Lake (blue) in northern Utah, USA. Right inset shows the northern tip of Antelope Island. Map baselayers are from the U.S. Geological Survey, National Geospatial Program ( https://basemap.nationalmap.gov/arcgis/rest/services/USGSTopo/MapServer and https://www.sciencebase.gov/catalog/item/52c78623e4b060b9ebca5be5 ). (B) Satellite images of Great Salt Lake from October 29, 2012 and (C) October 28, 2022 showing the shrinking shoreline of Great Salt Lake (MODIS corrected reflectance images from NASA Worldview). (D) Map of Great Salt Lake (at 1280 m elevation) showing the approximate location and extent of submerged vs. exposed microbialite reef areas in summer 2022, after Baskin et al. [ 13 ] . (E) Lake hydrograph from 1848 to 2022; area highlighted in gray is expanded in (F). The dashed gray line in both figures shows the historical (1963) lake lowstand. (G) Detail of field sites at Buffalo Point, with logger sites as vertical bars (B and B3), recovery experiment sites as horizontal bars (RA–RC), and the microbialite monitoring quad with monitored (M1–M3) and cored (C1–C3) microbialites. The 2020 shoreline is also shown as a dashed line. The underlying aerial view (from Google Earth) shows the site in May 2022, with bright areas showing exposed, desiccated microbialites.

Saline lakes around the world are facing a “desiccation crisis”: threatened by water overuse and climate change, with wide-ranging consequences to regional and hemispheric ecosystems, air quality, weather patterns, economic activities, and more [ 1 – 3 ]. In this paper, we present a case study of the largest terminal lake in the Western hemisphere: Great Salt Lake, in northern Utah within the arid Great Basin, which has experienced a sustained decline in lake level driven by human overconsumption in its watershed ( Fig 1 ) [ 1 ].

Materials and methods

Field sites, time series data logging, and sample collection The work described in this study focused on a microbialite reef on the northern end of Antelope Island in Great Salt Lake (Fig 1). GPS coordinates of all measurement and sample locations are provided in S1 Table. Time series water pressure, temperature, and downwelling irradiance were measured every 15 minutes collected using data loggers (temperature/pressure: HOBO U20L, light/temperature: HOBO MX2202, Onset Computer Corporation) attached to a PVC pipe anchored to the lake bed. In March 2019, the instrument site was placed in 0.9 m deep water in a microbialite reef ~75 m from that date’s shoreline (Site B; Fig 1G). In August 2021, Site B became subaerially exposed (Fig 3A) and the logger pipe was moved to a deeper site ~150 m farther lakeward from the 2019 shore (Site B3; Fig 1G). In addition, manual water depth, visibility, salinity (using a handheld 0–28% refractometer with automatic temperature compensation; measurements are reported as a % by mass), density (using a brewing hydrometer), and temperature (using a digital aquarium thermometer) measurements were collected monthly to seasonally, along with microbialite surface observations. Lake elevation data were obtained from two U.S. Geological Survey monitoring sites in the lake’s South Arm (Fig 1A) in order to provide a continuous record of lake elevation during the period of our study: one near Saltair (ES1: Station 10010000, the standard site for Great Salt Lake south arm elevation measurements, which had an interruption in data collection from 2022-09-28 to 2022-12-14 due to historic lake level fall), and the other on the railroad causeway (ES2: Station 10010024, which operated from 2020-06-08 onwards) [38]. Multiple manual field measurements of site water depths at each site were then used to determine depth offsets vs. lake level (i.e., site elevation). During dates when both ES1 and ES2 recorded data, daily mean values at the two sites were averaged for inferring water depth changes at our field sites.

Weather data Weather data for 2019–November 2020 was obtained from a station on Antelope Island, located 4 km from the field site and operated by Antelope Island State Park (WS1: KUTSYRAC22, Ambient Weather WS-2090; Fig 1A); the station was non-operational beginning in November, 2020. Data for nearby stations available on WeatherUnderground (wunderground.com) were analyzed to find a new station with values consistent with those measured at KUTSYRAC22; for the full analysis see the file in the Open Science Framework data archive for this study (hereafter referred to as OSF archive). The station with the best coverage and closest similarity to KUTSYRAC22 was determined to be a private station located 14 km from the field site (WS2: KUTSYRAC27, Ambient Weather WS-2902; Fig 1A), with publicly available data retrieved and used in this study with permission from the station owner. For analytical purposes, measured weather values were averaged when data from both sites were available.

Microbialite field monitoring & core sampling In addition to general observations collected during the long-term monitoring work, detailed systematic monitoring of microbialites at the study sites was conducted from July 27–August 17, 2021 and July 12–August 2, 2022 as part of the Weber State University GETUP (Geoscience Education Targeting Underrepresented Populations) Summer Research Experience program. Microbialites monitored in summer 2022 were additionally visited and sampled sporadically until October 20, 2022. For this work, microbialites were flagged for repeat photography and sampling. In 2022, monitored microbialites (in addition to logger Site B3 and recovery experiment Site RC) were located within a roped-off rectangle (quad) to protect them from foot traffic, as the lake’s microbialites are currently unprotected. Each flagged microbialite was photographed during each visit, and the location of different colored bands on the surface of each microbialite were measured using a homemade surveying device (Fig S1.1 in S1 Appendix): the vertical distance of the band from the water surface, corrected for fluctuating lake elevation, was used to compare band height across dates. The vertical distance from the sediment/water interface was also measured, however the soft and mobile nature of the sediment surrounding the microbialites made measurement from the water surface the more reliable measurement. In 2022, core samples from microbialite tops were collected using 50 mL syringes with the tapered tip cut off, producing a coring tube that could be pushed by hand directly into the incompletely-lithified surfaces of the microbialites, to a depth of up to 4–7 cm. Cores were then extracted using the syringe plunger onto core cradles with a scale, and photographed. The 3–7 cm cores were sectioned in the field into three roughly equal 1–2 cm top, middle, and bottom (deep) sections using a sterile scalpel, then stored in sterile 15 mL centrifuge tubes on ice for transport to the lab. For downstream analyses, collected sections were coded based on their original depth within the microbialite as belonging to one of several horizons, with horizon T capturing the uppermost 0–1 cm, horizon M capturing 1–2.5 cm, and horizon B capturing 2.5–4 cm (see S1 Appendix for details). Back in the lab, core subsections were ground to a paste with a sterilized mortar & pestle to homogenize, then aliquoted for microscopy and chlorophyll extraction as described below.

Recovery experiments Recovery experiments involved submerging pieces of a desiccated microbialite back into lakewater, incubating for varying lengths of time, and recovering them for measurements of community regrowth. The desiccated microbialite (t 0 control) was collected in October 2016 from the beach at Ladyfinger Point (Fig 1A) at roughly 1278.6 m elevation, indicating that it had been subaerially exposed for at least two years at the date of collection, after which it sat undisturbed and dry on a laboratory windowsill until its use in this experiment. The t 0 control microbialite was broken into 1–7 g pieces, which were placed into individual nylon mesh bags (mesh size = 350 μm) that were attached to submerged PVC anchors (Fig S1.3 in S1 Appendix) such that the samples hung suspended in the middle of the water column at the start of the experiment. In 2021 (September 27–November 12), experiments were run at two sites: Site RA (near the edge of a microbialite reef with significant water movement), and Site RB (in the middle of a reef surrounded by microbialites with healthy periphyton), both of which had water depths to sediment of ~ 20 cm, and samples were suspended at ~ 10 cm beneath the water surface. Triplicate samples were collected at timepoints from 10–40 days. The experiment was repeated in 2022 (September 27–November 12) at Site RC (adjacent to logger instrument Site B3 in the middle of a different, initially healthy microbialite reef), at a water depth to sediment of ~ 40 cm, and samples were suspended at ~ 30 cm beneath the water surface. Triplicate samples at Site RC were collected at timepoints from 0–100 days. All three sites were hydrologically connected (open) to south arm lake water at the beginning of the experiments, however, Site RC became hydrologically closed during the course of the experiment due to rapid lake level decline. In 2021, samples collected at each timepoint were subsampled in the field for pigment and DNA extractions. Pigment subsamples were collected with surrounding lakewater in sterile 1.5 mL centrifuge tubes wrapped in electrical tape to minimize light exposure. DNA subsamples were collected by first swabbing an area of each sample and mixing the swab in DNeasy PowerSoil kit (Qiagen, Cat. # 12888–50) bead tubes that had been prepared with kit lysis solution. Then, a ~ 5 mm solid piece of each sample was also broken off using sterile tweezers and added to the same bead tube. Both pigment and DNA subsamples were flash-frozen on dry ice and stored frozen for transport. Back in the lab, DNA samples were thawed, vortexed for 5 minutes, and stored frozen at -20°C until extracted following kit protocols (see S1 Appendix for additional details). Pigment samples were processed immediately following the protocols described below. In 2022, instead of processing samples in the field, samples were stored in sterile centrifuge tubes placed on ice for transport, and processed in the lab. Whole samples were then ground and processed following the pigment extraction and microscopy protocols in the same manner described for the core samples, and DNA extractions were performed using the same protocol as for the 2021 recovery experiment samples.

Laboratory desiccation experiments A submerged microbialite was collected near Site B3 in July, 2022, then placed in an incubator at 30°C with one full-spectrum LED lamp (24W 3500 K full-spectrum lamp, Juhefa, Cat. # B08S7VSX6) and one UVA/UVB CFL lamp (23W 6500 K UVA/UVB lamp, Lucky Herp, Cat. # B082DYBQLL); spectra are shown in Fig S1.4 in S1 Appendix. The microbialite was allowed to desiccate for several days to weeks, and rinsed with distilled water at intervals between 6–52 days to simulate rain events. Microbialite surface coloration was measured immediately before and after rinsing events and periodically thereafter. Photographs were taken alongside a color card under standardized light conditions in order to measure surface coloration using the method described below. In addition, the dry mass of the microbialite was measured to assess removal of surface carbonate (weathering) during rinsing events.

Color analysis Microbialite surface coloration—specifically, the relative amount of green—was measured as an indicator of surface pigmentation for field microbialites and the microbialite in the laboratory desiccation experiment, with lack of green indicating surface bleaching. To quantify coloration, photographs were taken of microbialite surfaces alongside a standard color card (Pixel Perfect 24-Color Standard Calibration Chart). Color thresholding was done using the public-domain image processing software ImageJ (https://imagej.nih.gov), and the thresholded images were then used to quantify the green pixels in an image, with a full protocol described in S1 Appendix.

Microscopy Core and recovery experiment samples collected for microscopy were weighed (~0.2 g dry mass), vortexed with 1 mL of 2% PBS-buffered formaldehyde to fix, and stored at 4°C. For both phase contrast and confocal laser scanning microscopy (CLSM), wet mount slides were prepared from ~60–100 μL of sample that had been vortexed to suspend solid material, using clear nail polish to seal coverslips to prevent water evaporation and salt precipitation. Brightfield and phase contrast microscopy photomicrographs were collected using an Accu-Scope EXC-500 with an Excelsis MPX-16C camera and CaptaVision software. For confocal laser scanning microscopy (CLSM), 4’,6-diamidino-2-phenylindole (DAPI, 1 μL of 1 μg⋅mL-1 stock in sterile, nuclease-free water; Thermo ScientificTM Cat. # 62248) and calcein (1 μL of 100 μg⋅mL-1 stock in sterile, nuclease-free water; Invitrogen Cat. # C481) fluorescent probes were added to the fixed sample prior to fixing cover slips, and fixed slides were stored in a dark box prior to analysis to prevent photobleaching. CLSM photomicrographs were imaged using an Olympus FV3000 with the following channels: DAPI (for staining DNA), ex = 405 nm, em = 430–480 nm; calcein (highlighting bound Ca2+ and Mg2+, which can be used to image polymers and biogenic carbonate), ex = 488 nm, em = 500–540 nm; and chlorophyll, ex = 514 nm, em = 550–600 nm (full settings used for imaging are in S1 Appendix). For both phase contrast microscopy and CLSM, ten random photomicrographs were collected at 200x magnification for relative color/fluorescence analysis, and interesting features were photographed at various magnification; protocols for color/fluorescence analysis are described in detail in S1 Appendix. The Euhalothece culture in Fig 2D was imaged using an Olympus BX51 microscope using a DP27 5 MP capture card mounted on a U-TV1XC adaptor; positive phase contrast and DIC were achieved by pairing an Olympus CX-PCD Phase Contrast condenser on Ph3 setting and Nomarsky DIC filter with a 100x objective.

Chlorophyll extractions Chlorophyll extraction protocols used in 2021 and 2022 differed slightly in response to changes in equipment availability. In 2021, pigment samples were extracted by grinding solid samples to a powder using a sterile mortar and pestle, weighing the powder, then extracting overnight in 5 mL chilled, 100% acetone at 4°C in 9 mL glass test tubes. The next day, the glass tubes were centrifuged inside 15 mL polypropylene centrifuge tubes for 5 minutes at 3000 g to sediment solids. The supernatant, containing polar pigments, was scanned in a 3.5 mL quartz cuvette (Vernier) from 500–800 nm at 1 nm spectral resolution and 0.1 s⋅λ-1 averaging using a UV-VIS spectrophotometer (Cary 60, Agilent). In 2022, solid samples were weighed, added to 4°C chilled 90% acetone in 1.5 mL opaque black polypropylene tubes (Argos Technologies), vortexed 5 seconds to mix, and polar pigments were extracted overnight at 4°C. Prior to measurement, samples were again vortexed, then centrifuged at 1000 g to sediment solids. Supernatant containing pigments was then scanned in a 400 μL quartz cuvette from 350–1000 nm at 1 nm spectral resolution and 0.1 s averaging using the UV-VIS spectrophotometer. Samples with dense concentrations of pigments (with peak absorbance readings > 2.0) were diluted with chilled 90% acetone prior to reading. Chlorophyll a concentrations were then quantified using the equation described in Ritchie (2008), correcting for dilutions and normalizing to extracted sample mass. Values are reported as mg extractable chlorophyll a per gram of dry microbialite sample (mg/g).

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