Minerals as Windows into Habitability on Lava Tube Basalts: A Biogeochemical Study at Lava Beds National Monument, CA

Abstract

Lava tubes on Earth provide unique hydrogeological niches for life to proliferate. Orbital observations of the Martian surface indicate the presence of lava tubes, which could hold the potential for extant life or the preservation of past life within a subsurface environment protected from harsh conditions or weathering at the surface. Secondary minerals in lava tubes form as a combination of abiotic and biotic processes. Microbes colonize the surfaces rich in these secondary minerals, and their actions induce further alteration of the mineral deposits and host basalts. We conducted a biogeochemical investigation of basaltic lava tubes in the Medicine Lake region of northern California by characterizing the compositional variations in secondary minerals, organic compounds, microbial communities, and the host rocks to better understand how their biogeochemical signatures could indicate habitability. We used methods applicable to landed Mars missions, including Raman spectroscopy, X-ray diffraction (XRD), Laser-Induced Breakdown Spectroscopy (LIBS), and gas chromatography–mass spectrometry (GC-MS), along with scanning electron microscopy (SEM) and metagenomic DNA/RNA sequencing. The main secondary minerals, amorphous silicates, and calcite, formed abiotically from the cave waters. Two types of gypsum, large euhedral grains with halites, and cryptocrystalline masses near microbial material, were observed in our samples, indicating different formation pathways. The cryptocrystalline gypsum, along with clay minerals, was associated with microbial materials and biomolecular signatures among weathered primary basalt minerals, suggesting that their formation was related to biologic processes. Some of the genes and pathways observed indicated a mix of metabolisms, including those involved in sulfur and nitrogen cycling. The spatial relationships of microbial material, Cu-enriched hematite in the host basalts, and genetic signatures indicative of metal cycling also pointed to localized Fe oxidation and mobilization of Cu by the microbial communities. Collectively these results affirm the availability of bio-essential elements supporting diverse microbial populations on lava tube basalts. Further work exploring these relationships in lava tubes is needed to unravel the intertwined nature of abiotic and biotic interactions and how that affects habitability in these environments on Earth and the potential for life on Mars.

1. Introduction

Lava tubes are subterranean structures that form in volcanic regimes on Earth as the result of an initial lava flow that cools at the surface while hot lava continues to flow below. Eventually, the top layer solidifies, and the flowing layer drains out, leaving behind a long tube or cave beneath the surface [1,2]. Over time, the basaltic rocks that form the lava tube are permeated with meteoric water or volcanic aerosols laden with reactive ions and trace metals [3,4]. The influx of water that seeps through the basalts results in the formation of mineral crusts on the rock surfaces and within cracks of the lava tube walls and ceilings, or thick powders on the floors. These deposits are often composed of sulfates and carbonates, amorphous silicates, and various phases of Fe-oxides [5,6,7]. The speciation of those minerals is dependent on the water and rock chemistry and interior conditions, such as temperature, humidity, and presence of aerosols [3].

The abiotically precipitated secondary mineral deposits and host basalts within lava tubes act as habitable environments by providing cation- and nutrient-rich substrates for microbial colonization as scavenged Ca²⁺, Na⁺, Mg²⁺, Fe²⁺/Fe³⁺, Cu²⁺, Mn²⁺, and CHNOPS elements from the rocks, waters, and degraded organic matter are utilized throughout the life cycles of those communities [8,9,10]. Trace metals common in lava tubes, such as Cu, are essential components of certain enzymes at low levels, but they can also be toxic to life if concentrations are too high [11,12,13]. Autolithotrophy—metabolisms that weather rocks to produce nutrients and energy—often serves as the central process in these oligotrophic environments [14,15]. While autolithotrophy may be dominant, a wide variety of metabolisms supporting diverse and unique microbial communities in these mostly dark and isolated environments have yet to be fully characterized [16,17,18,19,20,21,22].

The release of organic acids by microbes is highly effective for weathering silicates, as this process decreases the pH and induces metal complexation [15]. In addition, microbial materials (e.g., extracellular polymeric substances, cells, sheaths) are made up of a variety of proteins, lipids, enzymes, etc., that are collectively negatively charged to attract the positively charged cations serving as templates for mineral formation [23,24]. The variations in biotic elemental cycling and subsequent mineral precipitation can leave behind measurable chemical and morphological signatures that are indicative of life [12,25,26]. Though not always easy to distinguish in natural samples, minerals that precipitate through biotic means can have observable characteristics that differ from those that are purely abiotic [27,28]. This becomes important for life detection efforts in environments where the preservation of biotically derived organic material is less likely to occur, and only the minerals remain [29].

To accurately assess what processes are occurring in the formation or alteration of minerals in terrestrial environments, multiple analytical techniques are typically needed [25,30,31]. Raman spectroscopy, a laser-based vibrational spectroscopy technique, has long been used in geochemical studies to characterize the organic and inorganic composition of natural geologic samples [5,32,33,34,35,36]. The main challenge using Raman spectroscopy in this context is the overlap of peaks between different components in the same spectrum, as well as natural fluorescence, both of which can obscure diagnostic information of a probed sample [37,38]. In such cases, X-ray diffraction (XRD), another well-established technique to identify minerals, is often useful in clarifying which mineral phases are present [28,39]. To better understand how microbes are associated with minerals, techniques like Laser-Induced Breakdown Spectroscopy (LIBS) and scanning electron microscopy (SEM) coupled with energy-dispersive X-ray spectroscopy (EDS) can help identify the elemental variations in the minerals and their relationships with microbial material [5,11,40]. Gas chromatography–mass spectrometry (GC-MS), a compound-specific organic chemistry technique, allows the separation and identification of a large range of compounds, and metagenomic DNA/RNA sequencing to assess general microorganism types and metabolisms further complements the above techniques [5,14,41,42].

Raman spectroscopy, XRD, LIBS, and GC-MS have all been employed by Mars rovers to search for signatures of life and to assess the habitability of the Martian surface [43,44,45,46]. Orbital observations of the Martian surface indicate the presence of surface structures that resemble surface features of lava tubes on Earth, including sinewy structures, collapse pits, and skylights [1,47]. If lava tubes exist on Mars, they would provide habitable, sheltered environments protected from the harsh conditions at the surface that could hold the potential for extant life as well as for the preservation of past life [48]. The lithologies of the rocks at Lava Beds National Monument, CA, are similar to those targeted by Mars rovers and provide an ideal field site to study the compositional variations in basalt and crust minerals, organic compounds, microbial communities, and how their biogeochemical signatures could be potential indicators of habitability [49,50].

We conducted a biogeochemical investigation of two basaltic lava tubes in the Medicine Lake region of the northern California Cascades by characterizing the compositional variations in secondary minerals, organic compounds, microbial communities, and the host rocks using a suite of instruments relevant to Mars missions. Our main goal was to understand how the mineralogy is shaped by microbial communities in basaltic lava tube systems and how some minerals may provide habitable environments for the microbes to thrive. A better grasp of their biogeochemical signatures will also be useful for life detection on rocky planetary bodies beyond Earth.

2. Materials and Methods

Samples of secondary precipitates (crusts) and the host basalts were collected from different locations inside two different lava tubes in Lava Beds National Monument (LBNM), CA: Golden Dome and Hercules Leg (Figure 1). All samples were collected from isolated areas not exposed to sunlight or the surface. Golden Dome (GD) is about 679 m long and is characterized by a golden sheen coating many of the ceilings and some of the walls, which has been attributed to the concentration of water droplets by Actinobacteria [19]. Water was observed mainly on one side of the tube, seeping down the walls or dripping from the ceiling, and there were a few areas on the floor with standing water (~ a few cm’s deep). Hercules Leg (HL) is about 594 m long and much drier in comparison to GD, with only a few areas where appreciable water was observed. The four sampling locations in HL included an interface of the ceiling and upper wall (HL1-T) and a lower wall near the floor (HL1-L), an area of white, brown, and beige crust on an upper wall (HL-2), and an orange crust on the wall (HL3). The four sampling locations in GD included a gray crust on an upper wall with beige-yellow and white-tipped spicules (GD-1), beige and yellow on gray crusts on a lower wall (GD-2), black coatings on beige crust on a low ceiling (GD-3), and pink and beige on gray spicular crust on an upper wall (GD-4). Samples were collected cleanly from both tubes using ashed tools and glass jars and were kept on ice until returned to the laboratory, where they were kept in cold storage (−30 °C and −80 °C) or in a desiccation cabinet (subsamples for mineral analyses). A subset of samples was cleanly collected for metagenomic analyses, stored and transported in liquid nitrogen, and kept in cold storage (−80 °C) in the laboratory. Images of the sampling locations are shown in Figure 1, and descriptions of each sample discussed here are summarized in

Table 1. The source areas, mineralogy, and elemental and molecular composition detected with Raman spectroscopy, XRD, EDS, and LIBS in lava tube samples.

Sample	Source Area	Mineralogy	Elemental Composition		Organic Compounds
		Raman & XRD	UV-VIS LIBS	EDS	Raman	LWIR LIBS
HL-1L	Chamber after 2nd entrance, lower left wall near floor	Amorphous silicates, smectite clays, calcite, plagioclase, Cu-hematite	Major: Ca, Ti, Al, Mg, Si, Na, Cu, V Minor: S, Fe, Mn, Ni, Cr, C2, Ti	Si, Al, Ca, Mg, Na, K, C, N, S, P, Cl, Fe, Ni, Ti	Organic acids, N-compounds	H2O, O-Si-O, SCN CO, CN, COC
HL-1T	Chamber after 2nd entrance, upper left wall above HL-1L	Amorphous silicates, smectite clays, calcite, gypsum, orthoclase, pyroxene, hematite, ilmenite, magnetite, Ni-ferrite, Cu-hematite	Major: Ca, Al, Mg, Na, V Minor: S, Si, Fe, Mn, Ni, Ti, Cu	Si, Al, Ca, S, Mg, Fe, C, Na, N, K, Ni, P, Cl, Cu, Ti	Organic acids, carotenoids	H2O, O-Si-O, O-S-O
HL-2	Chamber after 2nd entrance, low ceiling	Amorphous silicates, gypsum, Cu-Fe-oxides, hematite, Ni-Ferrite, pyroxene, anorthite	Major: Ca, Fe, Mg, Si, Na, Cu, V Minor: S, Ti, Al, Mn, Ni, Cr, C2	Si, Al, Ca, S, Fe, Mg, C, N, Cu, Ti, P, Cl	Organic acids, carotenoids, N-compounds	H2O, O-Si-O, SCN CO, CH, CN, COC
HL-3	Distant Light; 1st entrance, around a wet opening	Amorphous silicates, smectite/vermiculite clays, Fe-oxides, anorthoclase	Major: Al, Fe, Ti, Mg, Si, Na, Cu, V, Ni, Cr Minor: S, Ca, C2	Si, Al, Ca, Fe, Mg, C, N, Ni, Zn, Ti, K, Na	Organic acids	H2O, O-Si-O, OCN, SCN CN, COC
GD1	Chamber 1, ceiling + wall	Amorphous silicates, smectite clays, calcite, anorthoclase, pyroxene, hematite	Major: Ca, Fe, Ti, Mg, Si, Al, Na, Cu, Li Minor: S, Cr, V, H	Si, Al, Mg, Ca, S, C, Na, Fe, K, N	Organic acids	O-Si-O, CN, COC H2O, O-S-O
GD-2	Chamber 1, wall, small ledge	Amorph silicates, Na-plagioclase, pyroxene, calcite, FeCu-oxide	Major: Mg, Si, Na, Ca, Cu Minor: S, Fe, Cr, C2, V, Li, Al, Ti	Si, Al, Mg, Fe, Ca, C, N, P, Na, K, Cu, Cr	Organic acids	H2O, O-Si-O, SCN CO, CN
GD-3	Chamber 1, ceiling + wall	Amorph Si, quartz, anorthoclase, Fe-oxide	Major: Ca, Mg, Si, Al, Na, Cu, Li Minor: S, Fe, Al, Cr, C2, V, Ti	Si, Al, Mg, Ca, C, S, Fe, N, Cu, K, Na, P, Cl, Cr, Ag	Carotenoids, N-compounds, organic acids	H2O, O-Si-O, SCN, CO, CN
GD-4	Chamber 1, opposite wall: low ceiling/wall	Amorph silicates, smectite clays, anorthoclase, Fe-oxides, calcite, clinopyroxene	Major: Ca, Mg, Si, Al, Na, Cu Minor: S, Fe, Li, Cr, V, Ti	Si, Al, Ca, Mg, Fe, Na, S, C, K, Ti, Cl, P	Organic acids	O-Si-O, CO, CN

.

Field measurements were performed using a portable VIS Raman spectrometer (TSI, Inc., EZ Raman, Shoreview, MN, USA), which uses a 532 nm excitation laser with a 1.2 μm fiber optic probe and a spot size of 50 µm. Calcite (CaCO₃), gypsum (CaSO₄·2H₂O), and quartz (SiO₂) minerals were used as calibration standards to keep track of any spectrometer drift and to accurately assign peak positions to our samples. The instrument has a spectral range between 100 and 4000 cm⁻¹, with a resolution of ~10 cm⁻¹. Power output is controlled by the software, and a minimum amount of power (0.3–2.0 mW) was applied as appropriate for each individual sample. Scan times varied for each sample, typically ~10–40 s/spot for each target, while some required up to 180 s/spot. Images of the probed areas were collected for context using a Dino-Lite handheld digital microscope (Dunwell tech, Inc., Los Angeles, CA, USA).

High-resolution micro-Raman measurements were performed on the samples that were collected and returned to the laboratory with a WITec α-Scanning Near-Field Optical Microscope (WITec GMBH, Ulm, Germany) customized to incorporate confocal VIS (488 nm and 532 nm) Raman spectroscopy imaging utilizing 50× and 100× objectives to achieve a lateral resolution of ~300 nm and a spot size of ~1 μm. Spectra maps and spot scans were performed using low power at the sample (~0.05–3 mW) with acquisition times ~1–2 s/pixel for each map and 3–120 s/per spot for spot scans.

The Raman data collected with the portable instrument were processed using ACD Labs Optical Workbench and Spectrus Processor software packages (2023.1.1). Baseline correction and peak fitting were performed manually using Gaussian–Lorentzian fits. For the micro-Raman imaging data, WITec Project Plus (V5) software was used to map peaks of interest across the sample and to compute peak intensity maps. Gaussian–Lorentzian fits were also used to baseline correct the spectra and to assign peaks. Interpretations of the Raman spectra were based on careful comparisons to publicly available databases (e.g., RRUFF), our previous work, and published literature (cited where relevant).

The mineral compositions of secondary precipitates and host rocks were also determined by X-ray diffraction (XRD) analyses. Sample fines were prepared by crushing the precipitate crusts to a fine powder via mortar and pestle or drilling material from the surfaces of samples using a Dremel (Mt. Prospect, IL, USA) rotary tool when necessary. XRD patterns were acquired from 2 to 70° 2Theta (Cu Kα radiation, k = 1.54059 Å), at 0.01°/step and at least 2 s/step using a Bruker D8 Discover diffractometer (Bruker, Billerica, MA, USA). Mineralogy was obtained using the MDI (Materials Data Incorporated) Jade analysis software v9.

Compositional and textural features of the samples were characterized using scanning electron microscopy (SEM) and energy-dispersive X-ray spectroscopy (EDS). Rock and crust fragments were mounted on carbon tape and analyzed uncoated with a Phenom ProX SEM-EDS (Thermo Fisher Scientific, Waltham, MA, USA) system for high magnification imaging and basic chemical composition of grain surfaces, microbial material, and mineral precipitates [27]. EDS analyses were performed at 10 kV or 15 kV with a spot size of <1 mm. The relative elemental composition was determined using Phenom ProSuite v2.9.0 software.

UV-VIS-NIR (UVN, 200–1000 nm) + Long-wave IR (LWIR, 5.6 to 10 µm, 1785 to 1000 cm⁻¹ in the mid-IR region) Laser-Induced Breakdown Spectroscopy (LIBS) (Brimrose Corporation of America, Sparks-Glencoe, MD, USA) was also performed on a subset of the samples to confirm elemental and molecular composition. Spectra were obtained by averaging results from four laser pulses sampling different spots on the sample surface. Calcite (CaCO₃), gypsum (CaSO₄ 2H₂O), quartz (SiO₂), magnetite (Fe₃O₄), alumina (Al₂O₃), feldspar and augite minerals, plus a basalt from a GSI 1050-00E mounted specimen set, were used as calibration standards. Fragments of the samples collected and returned from the field were mounted vertically on an XYZ translational sample stage that was positioned 1 m away from the front end of the collection reflecting telescope and probed in the ambient atmosphere. Broad spectra of emission signatures in both the UVN and LWIR spectral regions initiated by the same single laser-pulse-induced micro-plasma on the target surface were recorded with a 3 µs delay with a 10 µs integration window for the UVN spectrometer and a 24 µs delay with a 44 µs integration window for the LWIR spectrometer [51]. Origin Pro 2023 spectroscopy software was used to analyze all LIBS data.

The organic composition of the samples was determined by pyrolysis GC-MS using a Frontier Laboratories 3030D multi-shot pyrolyzer (Quantum Analytics, Woodlands, TX, USA), and a Trace 1310 GC coupled to a quadrupole mass spectrometer (ThermoFisher, Waltham, MA, USA) equipped with a Restek (Centre County, PA, USA) capillary MXT-5 column (30 m long × 0.25 mm ID bonded with a 0.25 µm thick stationary phase). Prior to analysis, samples were powdered using solvent-washed agate mortar and pestle and ashed tools, and aliquots (~10 mg) were deposited into the pyrolysis stainless-steel cup. The samples were pyrolyzed using a slow pyrolysis ramp of 35 °C·min⁻¹ within two temperature ranges: first, 55–250 °C to remove any adsorbed water or volatiles, then 250–950 °C intended to extract the organic molecules and potential biosignatures trapped in the minerals that decompose within this range (e.g., clays, sulfates, carbonates). During pyrolysis, volatilized molecules were trapped at the column inlet using a liquid nitrogen cold trap (−180 °C); the GC injector downstream was set between 250 and 300 °C to prevent condensation. After pyrolysis, the cold trap was heated, automatically triggering the beginning of the GC analysis. The temperature program used for the GC was an initial temperature of 35 °C held for 5 min, followed by a 5 °C.min⁻¹ ramp up to a temperature of 200 °C, followed by a second 10 °C.min⁻¹ ramp up to a final temperature of 320 °C held for 1 min for bakeout. Helium was used as the carrier gas and set at a 1.2 mL.min⁻¹ flow rate with splitless analyses or 10–30 split ratios depending on the sample. The MS ion source and transfer line were both set at 300 °C, and ions produced by the 70 eV electron ionization source were scanned with mass-to-charge ratios (m/z) between 40 and 550 with a scan time of 0.2 s. Flight-like wet chemistry experiments, and more specifically, tetramethylammonium hydroxide (TMAH) thermochemolysis, were also performed, analogous to one of the wet chemistry techniques currently utilized onboard the Sample Analysis at Mars (SAM) instrument of the NASA Curiosity rover [45].

For DNA extraction, samples from Golden Dome and Hercules Leg caves were extracted in duplicate and underwent a pre-extraction chemical lysis in which ~250 mg of sample was mixed with 400 µL 2X PBS, 25 µL metapolyzyme (Millipore Sigma, Milwaukee, WI, USA, cat. no. MAC4LDF-5X1VL), and 250 µL ZymoBIOMICS DNA Miniprep Kit Lysis Solution (ZymoBIOMICs, Orange, CA, USA). Each mixture was initially vortexed for 10 s and incubated at 35 °C for 2 h, with intervals of 10 s of vortexing every 30 min. An amount of 5 µL of proteinase K was then added to each sample, followed by another incubation at 55 °C for 1 h. The lysed samples were then processed using steps 4–11 of the ZymoBIOMICS DNA Miniprep Kit (ZymoBIOMICs, Orange, CA, USA cat. no. D6035). Paired-end 150 bp libraries were prepared using the Nextera XT DNA Library Preparation Kit with Illumina DNA/RNA UD Indexes (Illumina, San Diego, CA, USA) and sequenced on a NovaSeq platform. Reads were trimmed to remove adapters and low-quality bases using Trimmomatic v0.39, assembled using SPAdes genome assembler v4.0.0 (metaSPAdes mode), and filtered for a minimum length of 500 bp using bbtools v39.08 [52,53]. Functional and taxonomic gene annotation was performed with JGI IMG/M ER v6.0.0 [54].

3. Results and Discussion

Amorphous silicates were identified as the predominant secondary mineral in crusts and coatings from both lava tubes by Raman spectroscopy, XRD, and SEM/EDS, which is in agreement with previous studies [55,56]. The amorphous silicates were identified with Raman spectroscopy by peaks ~336 cm⁻¹ (O-Si-O deformation), 398 cm⁻¹ (Si-O-Si bending), 449–480 cm⁻¹ (Si-O-Si), 538 cm⁻¹ (Si-O-Si bending + M), 730–772 cm⁻¹ (Si-O-Si symmetric stretch), 930 cm⁻¹ (Si-O stretch), and 1044 cm⁻¹ (Si-O-Si asymmetric stretch) [57,58]. The maxima for the broad features in all Raman spectra were centered between 410 cm⁻¹ and 480 cm⁻¹, suggesting that the main phase is opal-A [59,60]. A broad “hump”, centered near a d-spacing of ~4 Å, was observed in the XRD patterns, which is typically attributed to silica-rich X-ray amorphous material similar to opal-A [60,61]. The SEM images show the pervasiveness of the amorphous silicates from both lava tubes, ranging from masses of silica spherules to layers of crusts up to 100 mm thick (Figure 2). EDS data typically mainly showed constituents Al and Mg in addition to the Si, and O in the silica-rich materials, and Ca, Na, and Fe were also observed less frequently (Figure 2).

The LIBS spectra from the silica-rich samples revealed a complex elemental composition with both major and minor constituents (Figure 2D). The most prominent emission features belong to silicon (Si), which appears strongly at 251, 288, 385, and 390 nm, and again near 634 nm, consistent with the dominance of silica in the sample. Magnesium (Mg) exhibits strong peaks at ~279.5 nm, 285.2 nm, and 517 nm, and sodium (Na) is clearly identified by its characteristic doublet at 589.0 and 589.6 nm. Calcium (Ca) is extensively present with intense lines at 393.4 nm, 396.8 nm, 422.7 nm, and several others at 616, 643, and 646 nm, indicating the presence of Ca-bearing phases. Copper (Cu) is visible with moderate-intensity peaks in the 320–330 nm range. Among the minor elements, sulfur (S) appears near 470 nm, iron (Fe) is distributed throughout the UV and visible regions, and chromium (Cr) shows a clear signal at ~263 nm. Additional minor species include vanadium (V), aluminum (Al), and titanium (Ti), primarily in the 300–340 nm region, and lithium (Li) with a weak but distinct peak at 670.8 nm. Carbon-related C₂ Swan bands spanning 467–473 nm suggest the presence of carbonaceous material or ambient molecular contributions [62]. A strong hydrogen (Hα) line at 656 nm and an oxygen (O) emission near 777 nm further point to atmospheric or moisture interactions during plasma formation. The high-intensity, well-resolved peaks and broad elemental coverage affirm the efficacy of LIBS in revealing both matrix and trace components in silica-rich amorphous samples.

As noted in other studies of lava tubes, our observations of composition indicate that water-basalt interactions are one of the primary mechanisms of amorphous silica formation in these lava tubes [56]. The uneven layers and patchiness of crusts we observed also likely reflect the fluctuation of evaporation and condensation within the lava tube over time [3]. Multiple microbial features appeared to be encrusted or replaced by amorphous silica as observed with SEM, including filaments and trichomes (individuals and masses), coccoids, tetrad clusters, and extracellular polymeric substances (EPS) (Figure 3). Coarse-grained amorphous silicate-rich and clay-rich areas were often interspersed with masses of microbial material as well. The inconsistent appearance (smooth vs. rough) of silica-coated microbial materials also suggests that the silica is passively formed on some materials and not on others due to the environmental conditions and not due to active biomineralization. EDS identified C (~1–11 Wt%), N (~1–12 Wt%), and P (~0.6–2 Wt%) in the microbial materials from both lava tubes that did not appear to be associated with carbonate or phosphate and could be attributed to organic compounds.

Hydrated silicates were also identified in both lava tubes by XRD, SEM/EDS, LIBS, and Raman spectroscopy (Table 1). Smectite clays were identified by Raman bands ~3575 cm⁻¹ (AlFeOH), 3440 cm⁻¹ (HOH), 3354 cm⁻¹ (HOH), and 3120 cm⁻¹ (HOH stretch) [63]. Na-poor vermiculites were identified less frequently by Raman bands ~3720 cm⁻¹ (Mg₃OH), 3450 cm⁻¹ (HOH), 3240 cm⁻¹ (Al-OH), 930 cm⁻¹ (Si-O), 754 cm⁻¹ (Si-O), and 410 cm⁻¹ (O-Mg-O). The higher frequency hydration Raman bands 3000 cm⁻¹–3800 cm⁻¹ were highly variable in all samples, reflecting a wide range of hydration states and phase transitions [63]. XRD did not indicate clay minerals present at abundances above XRD detection limits in our bulk powder samples, but it did show broad features indicating a high frequency of X-ray amorphous materials (likely Si-rich) in both lava tubes. The EDS data showed high Mg and Al with lesser Ca, Na, and Fe content in many amorphous silica-rich coatings and crusts observed with SEM. The LIBS data were mostly in agreement with EDS, but with strong signatures for Ca and Mg detected in all of the samples.

Plagioclase feldspar and pyroxene were identified as the main basalt minerals by Raman spectroscopy and XRD. Both minerals are expected to weather to poorly crystallized material and then smectite in the presence of water and under the ambient temperatures of the lava tube environments, though their dissolution rates are highest under acidic (pH 4–6) conditions [64,65]. Kulkarni et al., 2022 reported the pH of ~7 for the cave waters at LBNM, and a neutral pH is typical for basaltic lava tube systems worldwide [5,19,56,66]. The formation of the clay minerals may be enhanced by microbial processes under the environmental conditions within the lava tubes we studied [67]. During the bioweathering of silicates, the organic acids and EPS released by microbes effectively remove cations and break bonds within the silicate lattice, providing nutrients for the microbes and altering the mineral substrate [68,69]. In the process, the local pH can be decreased, creating conditions more conducive to secondary mineral formation [15,64]. SEM images show microbial filaments on or near clay-like textures among flat pyroxene or plagioclase surfaces in some of our samples (Figure 3). The spatial relationships between the swathes of microbial material and organic compounds with regions where the clay minerals/mineraloids formed suggest that the formation of the clay minerals we observed could be due to the microbial activity in those areas.

Samples from GD exhibited the most variety of microbial features, including filaments and EPS with an abundance of coccoidal forms, while the HL samples seemed to lack coccoids and tetrads and instead contained an abundance of filamentous morphotypes and EPS (Figure 3). Evidence of microbial populations was most easily observed on the basalts just under or near the silicified layers, where distinct morphologies were still intact (Figure 4 and Figure 5). The Raman signatures for organic acids were identified by weak bands at ~696 cm⁻¹, 999 cm⁻¹, 1185 cm⁻¹, 1280 cm⁻¹, 1349 cm⁻¹, and 1602 cm^−1, and broad features between ~2800 cm⁻¹ and 3500 cm⁻¹ in almost all the silica-encrusted samples. Carotenoid pigments were detected by Raman bands ~1002 cm⁻¹, 1152 cm⁻¹, and 1506 cm⁻¹ along with nitrogen-bearing compounds (M-CN) ~2148 cm⁻¹, (N=C=O) 2289 cm⁻¹, and S-bearing compounds (S-H) 2643 cm⁻¹ (Figure 4). Multiple biomolecular signatures were detected with LWIR LIBS in the crusts, including emission bands for CO, SCN, CN, COC, and CH (Table 1, Figure 4). The Raman and LIBS signatures indicating N-bearing compounds observed in the HL and GD samples are similar to those seen in a lava tube on Mauna Loa, Hawaii, and may indicate an active microbial N-cycle in lava tube environments [5,70,71].

The taxonomic groups we observed in our samples are similar to those in other basaltic lava tube systems, where similar secondary minerals (e.g., amorphous silica, gypsum, calcite) form and reflect diverse potential lifestyles present in the community, including obligate aerobes, chemoheterotrophs, and nitrogen fixers [14,21,22]. The metagenomic results indicated that samples from both lava tubes largely contained Pseudomonadota and Actinomyceota, with smaller amounts of Chloroflexota, Bacillota, Bacteroidota, and Acidobacteria. The Pseudomonadota were present as Gammaproteobacteria (Pseudomonadales, Lysobacterales), Alphaproteobacteria (Hyphomicrobiales, Sphingomonadales, Rhodobacterales), and Betaproteobacteria (Burkholderiales), while the Actinomyceota present are largely members of the Actinomycetes (Micrococcales) class. Some archaeal and eukaryotic sequences were present as Nitrosphaerota, Ascomycota, and Basidiomycota, respectively. The genes and pathways observed indicate a dominance of aerobic respiration via the tricarboxylic acid (TCA) cycle and the Entner–Doudoroff pathway, with autotrophy, photosynthesis, anaerobic respiration, and sulfate and nitrate pathways secondarily present. In addition, a previously unknown member of the Rubrobacteraceae was also identified in some of the GD samples in this study, which represents a facultative anaerobic heterotroph and potential methanotroph [72].

Gypsum (CaSO₄·2H₂O) is a commonly occurring precipitate from cave waters in most basaltic lava tube systems; however, it is not known to be a major component in the LBNM caves [55,56]. Kulkarni et al., 2022 concluded that the scarcity of sulfate minerals resulted from the low concentrations of S (~4 mg/L) and SO₄ (~1 mg/L) in the cave waters, with up to 80% of the sulfur in the organic form [56]. SEM imaging revealed large, euhedral grains of CaSO₄ with halite in areas where no microbial material or extensive silica had formed, only in the GD samples, indicating precipitation from cave waters. SEM imaging and EDS spectra also revealed the co-occurrence of cryptocrystalline grains of CaSO₄ spatially associated with silicified microbial material and masses of mineralized filaments or coccoids in samples from both lava tubes in areas where the host basalt surface intersected layers of amorphous silica (Figure 4 and Figure 6). These spatial associations between the microbial material and the cryptocrystalline gypsum suggest a second formation pathway involving the nucleation of sulfate on the cellular materials [73]. Gypsum was only rarely identified in the samples by XRD and by SO₄ Raman bands ~415 cm⁻¹, 493 cm⁻¹, 605 cm⁻¹, 620 cm⁻¹, 664 cm⁻¹, 1004 cm⁻¹, 1109 cm⁻¹, and 1138 cm⁻¹ in some of the HL samples [74]. It is possible that the gypsum was not detected with Raman more often due to its cryptocrystalline nature in most of the samples and its location within the areas dominated by amorphous silica.

A complex set of broad molecular vibrational features characteristic of silicate- and sulfate-bearing minerals, along with distinct signatures from water and organic species, was observed with LIBS (Figure 4). A prominent, overlapping band around 9.2–11 μm (1086–909 cm⁻¹) corresponds to Si–O–Si asymmetric stretching, while a similarly broad and intense feature at 8.9 μm (1123 cm⁻¹) is attributed to the SO₄²⁻ asymmetric stretching vibration [40]. The overlap of these two strong bands reflects the coexistence of amorphous silica and sulfate components within the matrix, likely forming a composite of layered mineral structure. In the 6.6–6.8 μm range (1470–1515 cm⁻¹), the spectra reveal pronounced water bending modes, along with a distinct vapor-phase H₂O feature at 6.6 μm (1515 cm⁻¹), strongly indicating the presence of structural or adsorbed water, as would be expected in gypsum-like hydrated sulfate minerals [75]. Additional emission peaks, such as the C=O stretching at 6.02 μm (1661 cm⁻¹), CN stretching at 8.4, 8.96, and 9.35 μm (1190, 1112, and 1069 cm⁻¹), and signals from isocyanate (OCN) and thiocyanate (SCN) at 6.98 μm (1433 cm⁻¹) and 9.05 μm (1105 cm⁻¹), highlight the presence of diverse organic sulfur-bearing and nitrogen-bearing compounds [76,77,78,79]. S-H bonds were also observed in Raman spectra of carotenoid pigments with gypsum, which correlates with the LIBS data and EDS data from the microbial materials, where enrichment in S (~1–6 Wt%) was observed (Figure 3).

Organic sulfur is a known component of microbial biomass that can be transformed and utilized in both the S- and N-cycles [80]. Dimethyl-sulfide (DMS), dimethyl-disulfide (DMDS), thiophene, and methyl-thiophene were detected in sulfate-bearing samples from HL with pyrolysis GC-MS. These S-bearing compounds can be derived from amino acids and readily cycled by microorganisms utilizing various metabolic pathways [81]. The metagenomic results indicated that samples from both lava tubes largely contained Pseudomonadota and Actinomyceota, with smaller amounts of Chloroflexota, Bacillota, Bacteroidota, and Acidobacteria. Several of these taxa could be involved in precipitating or inducing precipitation of the sulfate present in this environment, or their consumption may be the reason that sulfate was not widely detected. For example, the Pseudomonadota present in our samples, specifically the Burkholderiales, have previously been detected in lava caves and associated with sulfate-reducing metabolisms and abiotically precipitated lava tube gypsum [14,82]. Additionally, Burkholderiales are capable of both oxidation and reduction of sulfur compounds [83]. Gammaproteobacteria, Alphaproteobacteria, and Bacillota are also capable of sulfur oxidation, as well as EPS and biofilm formation, which have been shown to promote mineral nucleation and gypsum formation [22,84]. Members of the Alphaproteobacteria Hyphomicrobiales group we detected are capable of degrading DMS and oxidizing it to CO₂ and forming compounds such as sulfide, formaldehyde, and thiosulfate, while members of the Sphingobacteria group have been associated with the assimilation of DMDS [85,86]. It should also be noted that though the microbial materials can template crystal formation, gypsum may also be growth-limited in the areas rich in microbes, as laboratory studies have shown the inhibition of gypsum crystal growth in the presence of carboxylic acids [87].

Half of the total number of organic molecules detected in all of the samples with pyrolysis GC-MS included mono- and polycyclic aromatic hydrocarbons (PAHs) and aliphatic saturated and unsaturated hydrocarbons such as n-alkanes and n-alkenes pairs (Figure 5). Nitrogen-bearing organic molecules represented the second most numerous chemical family (15%–20% of the total number of organics), mostly in the form of aromatics: pyrroles, imidazoles, indoles, pyrimidines, benzonitriles (11%–17%), and fewer aliphatic compounds such as amines and nitriles (3%–4%), which correlates with the detection of N-bearing compounds with Raman spectroscopy and LIBS (Figure 3). Oxygen-bearing compounds were the second most numerous chemical family (~12% of the total number of organics) and were also detected as aromatics in the form of furans, phenols, naphthalenones, benzaldehydes, and aliphatics such as alcohols and ketones. Other aromatic forms of oxygen- and nitrogen-bearing molecules, such as pyridinones and quinolinols, were detected as well. A few chlorine-bearing molecules, such as chloromethane and C11-C15 chlorobenzenes, were also detected. We note that less than 10% of the molecules present in the chromatograms were not identified because of multiple coelutions and/or low match with the NIST mass spectra library [88].

TMAH thermochemolysis of the samples from both lava tubes yielded complex chromatograms with a high number of coelutions, preventing complete molecular characterization. Small sulfur-bearing compounds that were detected in the pyrolysis experiments could not be assessed due to coelution with solvent and TMAH decomposition products. Long-chain alkane/alkene pairs (C14–C32) were detected in HL-1T and HL-1L and in lower abundance (C16–C24) in GD-3. Detected molecular families included mono- and polycyclic aromatic hydrocarbons (PAHs), oxygen-bearing compounds such as furans and phenol esters, and nitrogen-bearing heterocycles such as pyrroles and indoles. HL-1T and HL-1L contained a higher relative abundance of PAHs in comparison to the other samples. Molecular biomarkers were also detected, including methylated sugars, fatty acid methyl esters (FAMEs), nucleobases, and amino acids. The detection of amino acids and nucleobases in particular is consistent with inputs from active microorganisms in the samples, as these compounds are degraded or recycled quickly in natural environments. Quantitative FAME content ranged from 150 to 3800 µg/g, with chain lengths between C11 and C18–C26. Nine to 46 distinct FAME species were identified in the samples, including linear, unsaturated, and branched.

Calcite was identified as a major component of the speleothems following amorphous silica in previous studies of these lava tubes [56]. In our study, calcite was detected in several of the samples by XRD (Table 1, Figure 2). Calcite was identified only weakly by Raman bands ~154 cm⁻¹, 282 cm⁻¹, and 1090 cm⁻¹ [89]. Several spectra collected from our samples exhibited high fluorescence, which could be caused by the calcite excited under the 532 nm Raman laser [90]. Evidence of calcite was also observed in the LIBS spectra by intense Ca lines at 393.4 nm, 396.8 nm, 422.7 nm, and several others at 616, 643, and 646 nm (Figure 2, Table 1). The SEM/EDS spectra did not inform associations between calcite and microbial material since both could contribute to any C detected with EDS, and no obvious morphologic or textural features indicative of calcite that could be spatially correlated with microbial features were observed with the SEM.

Hematite (Fe₂O₃) was also identified based on Raman peaks ~220 cm⁻¹, 293 cm⁻¹, 407 cm⁻¹, 660 cm⁻¹, and 1310 cm⁻¹. Some of the hematites were also mixed with other Fe-oxide phases, such as lepidocrocite based on peaks ~250 cm⁻¹, 530 cm⁻¹, and 650 cm⁻¹; magnetite based on peaks ~310 cm⁻¹, 380 cm⁻¹ and 670 cm⁻¹; wuestite based on peaks ~360 cm⁻¹ and 590 cm⁻¹; and goethite based on peaks ~478 cm⁻¹, 547 cm⁻¹, and 680 cm⁻¹ [91]. Some Fe-oxide phases, such as lepidocrocite, may not be expected to persist in these settings, but they can form from nitrate-reducing, Fe-oxidizing bacteria [92]. Large Fe-oxide rhombs (20–60 mm) were also observed on the basalt in zones rich in plagioclase, pyroxene, and smectite along the transition areas in the HL samples with SEM and EDS (Figure 6). Aggregates of small grains of Fe-oxides were observed in the GD samples in contrast to the large Fe-oxide rhombs observed in the HL samples (Figure 7). Dominant atomic emission lines from Ca, Fe, Mg, Na, and Si, characteristic of basaltic material, were observed with UV-VIS LIBS [93]. Calcium lines are especially intense at 393.4 nm and 396.8 nm (Ca II) and at 422.7 nm (Ca I), signifying a calcium-rich matrix. Multiple Fe emission lines are distributed throughout the 240–570 nm range, with notable peaks at 275.6, 358.1, 373.5, 404.5, 438.3, and 492.0 nm, indicating both Fe I and Fe II transitions (Figure 5). Magnesium is evident at 279.5 nm, 383.2 nm, and 517.2 nm, while sodium shows its characteristic doublet at 589.0 and 589.6 nm. Other elements detected in the host basalts included Si (~252, 288, 390 nm), Al (309.3, 396.2 nm), Cr (~263, 425 nm), Ti (~336 nm), Cu (~324.7 nm), and V (~318 nm).

Masses of microbial material were observed with SEM in the transition areas between silicate crusts and the underlying basalt surfaces, often in close association with the Cu-enriched (~2–8 Wt%) Fe-oxides (Figure 6). In their study of lava tube materials, Northrup et al., 2011 described remnant hematite rhombs associated with basaltic glass that were similar in size and at various stages of degradation [17]. Similarly to what we observed in our samples, masses of filaments were also observed between and on the rhombs in that study, suggesting that the rhombs formed abiotically and were later colonized by microbial communities [17]. The large size of the Fe-oxide rhombs in our samples facilitated correlated Raman measurements of the same grains (Figure 6). On the same grains characterized with SEM, the microbial filaments stretched across the rhombs had the distinct bright blue color typically attributed to Cu-rich material under the Raman microscope (Figure 6). FeCuO₃ and Cu incorporated with the Fe₂O₃ were identified by Raman peaks ~280 cm⁻¹, 340 cm⁻¹, 662 cm⁻¹, and a strong peak ~707 cm⁻¹ [94]. Biomolecular Raman signatures were identified with these Cu-rich phases, including carotenoids, along with bands ~1057 cm⁻¹, 1172 cm⁻¹, 1212 cm⁻¹, and 1548 cm⁻¹. Carotenoids were observed by Kopacz et al. in Cu-enriched silicates from an Icelandic lava tube [11].

A broad set of Raman bands ~1268 cm⁻¹, 1289 cm⁻¹ (-COOH), 1330 cm⁻¹ (CH₂), 1350–1362 cm⁻¹ (CH), 1380 cm⁻¹ (-COOH), and 1423 cm⁻¹ (-COOH) were observed either in place of or along with the 1310 cm⁻¹ hematite band (Figure 7). These compounds could be components of the EPS observed in these samples with SEM, as EPS is known to have a high binding capacity for Cu and Fe salts [95,96]. Organic acids were also observed in close association with Fe-oxides, plagioclase, or pyroxene of the basalts by Raman bands ~1111 cm⁻¹ (C-C), 1448–1460 cm⁻¹ (CH₂/CH₃), 2850 cm⁻¹ (_nsymCH₂), 2880 cm⁻¹ (_nasymCH₂), and 2940 cm⁻¹ (_nasymCH₃) [97,98]. Raman maps exhibiting the spatial relationships between these compounds and Cu-enriched Fe-oxides, combined with SEM images showing microbes with the same mineral phases, suggest a link between the microbes and secondary mineral phases, but the exact mechanisms cannot be determined based on our data (Figure 6).

The concentration of Cu in the cave waters is known to be ~23 μg/L, and Cu is an essential enzyme metal for some bacterial metabolisms [56,99]. Some of the taxa present in our samples could be involved in iron metabolism and/or promotion of copper mobilization. For example, the Actinomyceota we observed are capable of oxidizing iron under specific concentrations of nitrate, and some Pseudomonadales and Burkholderiales are known to produce siderophores for enhanced iron solubilization and copper mobilization from mineral matrices [100,101,102,103]. Weathering experiments have also shown that actinomycetes produced organic acids to facilitate both the leaching and complexation of metals from host rocks, with a strong preference for Cu [104]. Another taxon we detected, Bacteroides, is a diverse and understudied group that often produces carotenoids and has also been associated with the circulation of metal elements in lava tube environments [21]. Further taxa-specific laboratory studies focused on the biomobilization of Cu and other metals are needed to better understand the role of microbes in these systems.

Molecular vibrational emissions tied to both inorganic and organic species, similar to some of the other samples without Cu or Fe-oxides, were observed in these samples with LWIR LIBS as well. Emission features around 5.6 μm (1786 cm⁻¹) correspond to C=O stretching vibrations, while a broad H₂O bending feature centered at 6.6 μm (1515 cm⁻¹) indicates the presence of molecular or bound water, likely from hydrated sulfate phases such as gypsum [75]. In addition to these features, two broad, intense, and partially overlapping bands represent key inorganic oxide signatures: the SO₄²⁻ asymmetric stretching vibration at 8.9 μm (1123 cm⁻¹), confirming the presence of sulfate minerals, and the broad Si–O–Si stretching mode of amorphous or crystalline silica at 9.2–11 μm (1086–909 cm⁻¹), indicative of SiO₂-rich phases [40]. Organic signatures are especially pronounced in the mid-region: CH deformation bands appear at 8.03–8.4 μm (1245–1190 cm⁻¹), typically associated with amino acids or chlorophyll-like compounds. The OCN (isocyanate) band is present at 6.98 μm (1433 cm⁻¹), while strong CN stretching bands are observed at 8.96 μm and 9.35 μm (1112 and 1069 cm⁻¹). A C–O–C ether-like stretching feature is also visible at 9.05 μm (1104 cm⁻¹). In addition, a prominent SCN (thiocyanate) band was observed at 11.2 μm (893 cm⁻¹), reflecting sulfur-containing organics or secondary atmospheric interactions during the LIBS analysis [5].

NiFe₂O₄ (Ni-ferrite) was also identified by Raman bands ~330 cm⁻¹, 484 cm⁻¹, 665 cm⁻¹, and 698 cm⁻¹ and NiO by the recurring band ~504 cm⁻¹ in some of the samples from both lava tubes [105,106]. SEM-EDS identified a few Ni-enriched (~2 Wt%) Fe-oxide rhombs, but microbial material was not observed in any of the Ni-rich areas. Metal-oxide grains containing Ag (~40 Wt%) and grains of an Fe-, Cr oxide were also identified with EDS in isolated silica-rich areas devoid of microbial material in samples from GD. LIBS emission features for Cu, Cr, and Ti were detected in samples from both lava tubes, but Ni and Mn were only detected in the HL samples (Table 1).

4. Conclusions

(1)

The predominant secondary minerals we identified in the crusts and coatings of the lava tube samples were amorphous silicates, derived from basalt-cave water interactions. These precipitates were enriched in Al, Mg, Ca, Na, and Fe with lesser amounts of Cu, Cr, and V. Much of the microbial material we observed was permineralized with amorphous silica, which was enriched in C, N, P, and S.

(2)

Cryptocrystalline gypsum was identified at the interface between the basalts and amorphous silicate crusts. Sulfate and nitrate metabolisms indicated by some of the genes and pathways we identified, along with the S-bearing organic compounds detected as pyrolysis products or observed from Raman and LIBS analyses, and the spatial relationships between the microbial materials and the gypsum collectively indicate an active S-cycle in both lava tubes. It is not clear, however, if the microbes are actively inducing the sulfate formation or if they are utilizing abiotically formed sulfates as an energy source.

(3)

Clay minerals were also identified at interfaces between the basalts and amorphous silica crusts with Raman, SEM/EDS, and LIBS, often in areas where the cryptocrystalline gypsum was identified. The spatial relationships between the microbial materials and the clays and altered textures on basalt minerals observed by SEM suggest that the formation of these clays may be related to changes in local pH induced by microbial activity. We note that clay minerals were not detected with XRD, highlighting the importance of combining complementary techniques.

(4)

Large Fe-oxide rhombs and aggregates of Fe-oxide grains at the interface between the basalts and amorphous silicate crusts exhibited evidence of microbial colonization. Many of these were Cu-bearing Fe-oxides that were associated with microbial material, and we also detected taxa involved in iron metabolism and/or promotion of copper mobilization. Our combined results suggest that the Cu-enriched Fe-oxides we detected may have resulted from localized Fe oxidation and mobilization of Cu by the microbial communities.

The availability of bio-essential elements from cave waters and mineral surfaces supports diverse microbial lifestyles in the lava tubes at LBNM. The evidence for habitability in the lava tubes was collected using Mars-relevant instrumentation and highlights the utility of combined analytical techniques in the exploration of such environments. Further work targeting the interface between secondary mineral crusts and the host basalts is needed to unravel the intertwined nature of abiotic and biotic interactions, how that affects habitability in these highly dynamic environments on Earth, and what that might mean for similar environments on Mars.