organic matter

Spectral and biological characteristics of microbial mats and mosses across Fryxell Basin, Taylor Valley, Antarctica (2018-2019)


An intensive field campaign was conducted during the 2018-2019 austral summer to assess spectral and biological characteristics of multiple microbial mat types, as well as mosses, across nine ephemeral glacial meltwater streams in the Fryxell Basin of Taylor Valley, located in the McMurdo Dry Valleys region of Antarctica. In addition to biological sample collection, hyperspectral visible/near-infrared (VNIR) measurements were conducted using an ASD FieldSpec4 spectrometer, corrected and averaged across each mat, moss, and soil type, and downsampled to the multispectral resolution of the WorldView-2 satellite. This package contains a detailed sample archive, associated images, biological characteristics that include ash free dry mass (AFDM), chlorophyll-a, and pigment concentrations, as well as hyperspectral and multispectral reflectance spectra. 

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Field Methods

During the 2018-2019 austral summer, an intensive field campaign was conducted across nine ephemeral glacial meltwater streams (Bowles Creek, Canada Stream, Commonwealth Stream, Crescent Stream, Huey Creek, Lost Seal Stream, McKnight Creek, a Relict Channel, and Von Guerard Stream) in the Fryxell Basin of Taylor Valley, located in the McMurdo Dry Valleys region of Antarctica. 196 hyperspectral VNIR reflectance spectra were collected of individual and mixed microbial mat and moss communities using an ASD hyperspectral FieldSpec4 spectroradiometer (Malvern Panalytical). Measurements were referenced against a white Spectralon calibration target every 20-30 minutes or when illumination conditions changed. For each spectral measurement, it was broadly noted whether these microbial communities were dry, wet, or inundated. 139 of the total spectra were collected centimeters above the mat, moss, soil, or water (if inundated) in direct sunlight, representing 100% of that biological community or geologic material. 51 of the total spectra were collected around 140 cm above the ground and taken within a 30 cm diameter plastic ring where the percentage of mat, moss, and soil were adjusted between 0 - 100%. Other soil measurements were collected within six 20 x 20 m plots. In each plot, 75 spectra were collected approximately five feet, nine inches above the ground and averaged to produce six different hyperspectral measurements also included in this database. These plots were established for an atmospheric correction technique used in Salvatore et al. 2021. All hyperspectral measurements collected in the field were first splice corrected and then corrected using white reference measurements. Hyperspectral measurements were then averaged for each mat, moss, or soil type and downsampled to the resolution of the WorldView-2 satellite using methods outlined in Salvatore et al. 2021, which uses the spectral bandpass information contained in Updike and Comp 2010. Downsampling involves multiplying the filter responses for each band of the WV2 sensors by the hyperspectral measurements at each wavelength and summing all these values before dividing them by the sum of the filter responses at each band.

Alongside approximately half of all hyperspectral measurements, samples of 209 individual and mixed microbial mat and moss community types were collected within nine streams. Mat/moss types were categorized into thirteen different groups (see "mat_id" in the "archive" metadata for codes and descriptions). Red mat and sandy orange mat were classified as being pink and rubbery (red mat) and tan with a large soil matrix (sandy orange mat), but the true diversity of these communities cannot fully be represented in these simplified categories. Microbial mat, moss, and sediment samples were collected using standard MCMLTER methods: briefly, a #13 (13 mm) brass cork borer (2.27 cm2 diameter) was inserted into the mat to the underlying sediment and transferred to a sterile Whirl-Pak bag and filled with water from the stream. Tweezers were used when needed to transfer samples from the cork borer to the bag. For sediment samples, a credit card sized area was sampled to a depth of 1 cm and transferred to a Whirl-Pak bag . Samples that were dry at the time of collection did not have stream water added and remained dry. Three samples were collected for subsequent analyses that included ash-free dry mass (AFDM), chlorophyll-a, and other pigment measurements. Samples for ash-free dry mass and chlorophyll were filtered in the lab, wrapped in foil, and stored in the freezer. Pigment samples were filtered, put into cryovials, and then placed in liquid N2 for storage.

Images were taken of microbial mats and mosses and include .jpg and .NEF file formats (.NEF files are Digital Hemispherical Photos taken following the NSF NEON protocol and have not been processed for further analyses).

The coordinates (in decimal degrees) for the top left and bottom right corners of each square plot are indicated below:

Plot 1 for Bowles Creek

  • Top left corner = 163.053355 -77.624338
  • Bottom right corner = 163.052458 -77.624139

Plot 2 for Bowles Creek

  • Top left corner = 163.054699 -77.623749
  • Bottom right corner = 163.053781 -77.623545

Plot 1 for McKnight Creek

  • Top left corner = 163.269950 -77.597359
  • Bottom right corner = 163.269018 -77.597152

Plot 2 for McKnight Creek

  • Top left corner = 163.277794 -77.595986
  • Bottom right corner = 163.276990 -77.595800

Plot 1 for Crescent Stream

  • Top left corner = 163.202502 -77.649683
  • Bottom right corner = 163.201635 -77.649490

Plot 2 for Crescent Stream

  • Top left corner = 163.205806 -77.649483
  • Bottom right corner = 163.204937 -77.649286

Plot 1 for Canada Stream

  • Top left corner = 163.072247 -77.614833
  • Bottom right corner = 163.071364 -77.614626

Plot 2 for Canada Stream

  • Top left corner = 163.047767 -77.613291
  • Bottom right corner = 163.046902 -77.613099

Plot 3 for Canada Stream

  • Top left corner = 163.041647 -77.616134
  • Bottom right corner = 163.040718 -77.615932

Plot 1 for a relict channel

  • Top left corner = 163.284574 -77.625897
  • Bottom right corner = 163.283711 -77.625703

Plot 2 for a relict channel

  • Top left corner = 163.288517 -77.628270
  • Bottom right corner = 163.287635 -77.628074

Laboratory Methods

Algal AFDM and chlorophyll-a measurements were conducted in the Crary Lab at McMurdo Station. For AFDM, wet mass of samples were measured and dried at 100°C for 24 hours. Dried samples were weighed to obtain the dry mass, followed by combustion in a muffle furnace at 450 °C for four and a half hours. Ash-free dry mass measurements were calculated by subtracting the ashed mass from the dry mass. Percent organic matter was calculated by dividing the ash-free dry mass by the dry mass and multiplying by 100.

Chlorophyll-a samples were analyzed using a fluorometer. In a dark room, samples (including filter) were transferred to vials and 15 mL of a 90% acetone solution were added to each vial. Vials were vortexed for 15 seconds and transferred to 4 °C to extract for 24 hours. The vials were vortexed twice during the extraction period. Samples were diluted in 90% acetone as needed to be within instrument calibration range and 4mL were transferred to a cuvette and inserted into a Turner Designs 10-AU field fluorometer (Kohler 2018). The fluorometer was calibrated using a stock chlorophyll-a solution at multiple solutions to create a standard curve. The relationship between fluorescence and the concentrations of standards were applied to the sample measurements to obtain a chlorophyll-a measurement. Final chlorophyll-a concentration was calculated by adjusting for the dilution, which varied for each sample. Concentrations of chlorophyll were calculated using the McMurdo Long Term Ecological Research (LTER) Stream Team methods (see Kohler 2018).

Samples for detailed pigment analysis were freeze-dried in the Crary Lab in cryovials for 24-48 hours before being removed and stored in opaque nalgene containers at -20 °C. Samples were shipped frozen to Virginia Tech (USA) for storage prior to analyses.

High performance liquid chromatography (HPLC) was performed on 53 pigment samples in the Paerl Laboratory at the University of North Carolina Chapel Hill’s Institute of Marine Science in Morehead City, NC. Samples were weighed, and ~0.25 g of each sample was placed in a falcon tube for extraction. 1 mL of 100% HPLC-grade acetone was added to each falcon tube sonicated for 15 sec using a Sonics Ultrasonic Disruptor with microtip. Falcon tubes were stored dark in aluminum foil at -20°C for 20 - 24 hours. After extraction, 0.5 mL of the supernatant was filtered into amber glass autosampler vials using a syringe driven filter, Millipex Millipore 0.45 µm PTFE. Samples were kept chilled at 4°C and 200 µl of extract was injected into an HPLC system (Shimadzu system controller model CBM-20A, solvent delivery module LC-20AB) with an in-line UV/Vis photodiode array spectrophotometer (Shimadzu SPD-M20A; Pinckney et al., 1996, Jeffrey et al., 1997, Pinckney et al., 1999). The binary gradient consisted of solvent A (80% methanol: 20% ammonium acetate) and solvent B (80% methanol: 20% acetone). The extracts’ absorbance was measured by scanning the range of 350 - 700 nm every 1.28 sec. The data were collected and analyzed using Shimadzu’s LabSolutions Lite software, where individual pigments were identified using a combination of peak retention time, absorbance spectrum shape/signature, maximum wavelength and the similarity match of the unknown pigment to a standard. A multi-point calibration curve was generated by injection volumes of known quantities of pure pigment standards (manufactured from DHI, Denmark) and then calculating the peak areas of those pigments. The peak areas were then used to calculate the slope (response factor) for each pigment. Response factors were pigments that we did not have reference standards for were calculated using the ratio of the absorbance coefficients of each pigment to its closest structurally related reference pigment (peridinin in the case of scytonemin and scytonemin-red), multiplying the known pigment’s response factor by that ratio. Then, pigments extracted from the samples are quantified by multiplying the peak areas of a chromatogram at specific wavelengths by the response factors. Specifically, while the absorption maxima varies for each of the pigments analyzed, the Paerl Lab had chosen 440 nm as the baseline for efficiently quantifying all relevant pigments peak areas. Scytonemin is the most concentrated pigment in most of our samples, so we chose to calculate this one based on its own absorption maximum baseline. The absorption maximum for scytonemin and scytonemin-red in acetone is documented as 384 nm (Garcia-Pichel and Castenholz, 1991) and was visually identified as 388 nm based on the HPLC measurements, so peak areas for scytonemin and scytonemin-red were calculated using this 388 nm baseline. For the mass concentrations, the sample mass was subsampled from the total mass, and for areal concentrations, total area of the sample was determined from the AFDM measurements and subsampled for these analyses.

Mass and areal concentrations were included in this data package and calculated as follows:

ug/g pigment = [peak area]*[response factor]/[inj. vol.]*[filt. extr. vol.)]/[sample mass]


ug/cm2 pigment = [peak area]*[response factor]/[inj.vol.]*[filt. extr .vol.]/[sample area]

Additional information: 

Funding for this work was provided by the National Science Foundation grants #OPP-1758224 and #OPP-1745053 to Mark Salvatore, a Graduate Research Fellowship award to Schuyler Borges, and #OPP-1637708 and #OPP-2224760 to the MCM LTER for assistance with data management.


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