Preprints.org logo
Preprint
Review

This version is not peer-reviewed.

The Unresolved Paradox of Viable Microorganisms in Ancient Halite

Submitted:

08 September 2026

Posted:

09 September 2026

You are already at the latest version

Abstract
Reports of viable microorganisms from ancient halite create an unusual test of the biological limits of cellular survival. Halite deposits can be tens to hundreds of millions of years old, and increasingly rigorous studies have used chemical surface sterilization, petrographic selection of primary crystals, and direct sampling of primary fluid inclusions to reduce the likelihood of modern contamination. Nevertheless, the geological age of a crystal or inclusion does not by itself establish the age of a recovered organism. The central problem is that both major interpretations remain difficult. Modern contamination becomes less satisfactory when microorganisms are recovered from deeply buried, carefully sterilized primary material, yet uninterrupted survival for geological intervals is difficult to reconcile with spontaneous DNA and protein damage, ionizing radiation, finite energy supplies, and observed loss of microbial viability in other long-term environments. A further problem is molecular: several putatively ancient isolates or sequences are remarkably similar to modern organisms despite host-rock ages of tens to hundreds of millions of years. Proposed explanations include contamination, later recrystallization or groundwater infiltration, extremely low-activity maintenance states, and underestimation of genomic divergence by conserved markers such as 16S rRNA. None currently explains all observations. Ancient halite therefore provides a natural extreme-case model for a broader biomedical question: how long can a cell preserve sufficient molecular integrity for recovery when metabolism, repair, and replacement of damaged components approach their lower limits?
Keywords: 
halite
;  
halophiles
;  
microbial longevity
;  
cellular dormancy
;  
DNA damage
;  
molecular repair
;  
fluid inclusions
Subject: 
Biology and Life Sciences  -   Life Sciences
Introduction and Background
In October 2000, the reported revival of a Bacillaceae isolate from a brine inclusion of approximately 250-million-year-old Permian halite attracted considerable attention [1]. However, its genes were very similar to those of modern relatives [2]. The antiquity of the isolate was subsequently challenged, with modern environmental contamination proposed as an alternative explanation. Later efforts to isolate microorganisms from ancient halite found more archaea than bacteria (Table 1). Whole genomes of “ancient” microbes have been sequenced, and their phylogenetics and biochemical features have been analyzed [3,4,5]. Survival mechanisms of bacteria and archaea at extreme energy limits have been proposed and investigated [6,7,8,9,10]. Kinetics of biochemical processes that affect the lifespan of microorganisms have been studied at higher temperatures and extrapolated to environmental conditions [11,12]. Here we review a century of work on the identification and characterization of halophilic microorganisms in ancient halite, the properties of their genetic material, and proposed mechanisms of long-term survival.
For readers outside paleomicrobiology, the key distinction is simple but essential: the age of the salt deposit, the age of a fluid inclusion in the halite, and the age of a microorganism recovered from that inclusion are three separate claims. Geological and petrographic evidence can strongly support the first two without proving the third. The paradox arises because later studies increasingly reduced obvious routes of contamination while experimental biology simultaneously identified severe limits on how long an unrepaired or minimally active cell should retain a recoverable genome, proteome, and energy supply. This review evaluates both sides of that problem rather than assuming either ancient survival or modern contamination at the outset.
Why Ancient Halite Is Unusually Resistant to Contamination
Halite formations are widespread in the continental crust and can be massive in size. For example, the Salado formation from which the 250-million-year Virgibacillus was isolated covers 150,000 km2 with a thickness of 300-600 m. The ionic NaCl crystals are so compact that halite formations can be used to store radioactive waste and hydrogen gas [13,14,15]. The interior of intact halite beds can be strongly isolated from surface-water contamination. Even though halite may recrystallize under pressure, structural disturbances are localized, self-healing [16], and can be easily identified [17]
Microbes reside in fluid inclusions of halite crystals [18]. Fluid inclusions are small droplets of fluid, often less than 20 µm, that become trapped within a crystal either during the initial formation of that crystal or from a recrystallization event that occurred due to microfractures at a later time period [19]. Fluid inclusions formed during the initial growth of the crystal are termed primary inclusions while those that occurred due to microfractures and recrystallizations at a later period are termed secondary inclusions.
The chemical composition of fluid inclusions in ancient halite is consistent with that of ancient seawater. Using a model based on steady-state mixing of river water with mid-ocean ridge hydrothermal brine, Spencer and Hardie [20] calculated ion compositions in seawater throughout geological times. Ion concentration fluctuations in ancient seawater derived from this model matched the observed geochemical alternations in nonskeletal limestones and potash evaporites during the past 600 million years [21]. Subsequently, Timofeeff et al. [22] analyzed ion chemistry of primary fluid inclusions in halite crystals from multiple evaporite deposits, all from the Cretaceous period, and found that early Cretaceous halite inclusions had lower Mg2+/Ca2+ ratios than those in late Cretaceous halite, supporting the Spencer-Hardie model and the primary, syn-depositional origin of the inclusions, although inclusion age alone does not establish the age of microorganisms detected within them. Other physicochemical analyses have constrained the geological age and history of halite and its inclusions from geological periods such as Cambrian, Ordovician, and Permian [23,24,25].
How Contamination Controls Improved Over Time
Pioneer Studies
In 1960, Reiser and Tasch reported finding diplococci in Permian halite crystals of the Carey Salt Mine in Hutchinson, Kansas and subsequently culturing a halophilic Gram-positive diplococcus from dissolved crystals [26,27]. The crystals were submerged in 70% alcohol for 15 seconds before being dissolved in hyperosmotic media. At about the same time, Dr. Heinz Dombrowski of Justus Liebig University in Giessen, Germany, isolated multiple species of bacteria from halite from mines and drill cores of the ancient Zechstein Sea deposits [28,29]. He took scrupulous care in his lab setup, procedures, and sample preparation to eliminate laboratory contamination. He selected large primary salt crystals with diameters above 6 cm which “came from perfectly undisturbed points in the middle of larger successions of rock salt”, suspended them with a wire, flamed all sides for a total of one minute, cut the wire, and dropped the crystal into liquid medium in a glass tube which was then sealed and incubated at 40°C. The treatment was tested and found to be efficient with halite crystals dipped in Pseudomonas aeruginosa. The medium was saturated with NaCl as the halite dissolved. His success rate was about 50%. He phenotypically identified one species as Bacillus circulans and another as a new species which he named Pseudomonas halocrenaea. Dombrowski also isolated bacteria from Precambrian (650 Myr), Silurian, and Devonian halite from North American samples. Dombrowski’s claims were so controversial at the time that, according to Sir Fred Hoyle, he lost his tenured position at Justus Liebig University. De Ley et al. [30] thoroughly analyzed the phenotypes and DNA composition of Dombrowski’s new Pseudomonas and found it indistinguishable from Pseudomonas aeruginosa. However, Bibo et al [31] validated Dombrowski’s work by isolating Bacillus and other bacteria from Zechstein halite.
Revival and Expansion
Norton et al. [32] collected samples from two different British mines, Winsford salt mine in Cheshire, England and Boulby potash mine in Cleveland, England. The Winsford salt mine had bedded deposits from the Triassic period from 195-225 million years ago and samples such as brine and recrystallized salt sludges from pools, wall efflorescence, and newly exposed surface crystals from tunnel walls by blasting were collected. The Boulby potash mine was located in a Permian formation from 225-270 million years ago and samples such as halite, potash, brine pools, and efflorescence were collected. The surface crystals were submerged in absolute ethanol for 4-5 hours for decontamination and transportation. Isolates taken from cultures of these samples were chemotaxonomically characterized by their lipid profiles and found to be comparable to several representative haloarchaeal genera including Halobacterium (Hbt.) salinarum, Hbt. saccharovorum, Haloarcula, and Halococcus (Hcc.). While these isolates were from a variety of sample sources and not specifically fluid trapped within halite crystals, the authors argued for their syn-sedimentary origin due to the lack of typical surface colonization by haloarchaea in North Europe and the rapid loss of viability of these isolates in media without at least 1.5 M NaCl.
In an Austrian salt mine near Bad Ischl, Denner et al. [33] isolated a strain of Halococcus from 225–280-million-year-old Permian rock salt. The halite samples had clay on the surface, but they were dipped in ethanol and flamed [34]. The authors named the species Hcc. salifodinae and the isolate BIpT. The isolate shared similar features with previously sampled Halococci in 16S rRNA sequences, polar lipid composition, and menaquinone content, but differed in pigmentation, whole-cell protein patterns, and the presence of unknown lipids, justifying its classification as a new species. The group also isolated a species whose total cell protein constitution and membrane ATPase was similar to Hbt. saccharovorum.
Stan-Lotter et al. [35] compared multiple Halococcus isolates from European halite beds including the Austrian BlpT, two other strains isolated in 1997 from the same mine as BIpT, the British Br3 isolated by Norton et al. in 1993, and a previously unpublished 1988 isolate from a salt mine in Berchtesgaden Germany known as BG2/Thorough analysis of these different isolates led to the conclusion that they all belonged to the same species of Hcc. salifodinae. The authors proposed that since these organisms were all isolated from rock salt of similar geological age in widespread locations across Europe, the species was an inhabitant of connected ancient hypersaline seas covering much of the European continent today.
Radax et al. [36] obtained halite crystals from mines and drill cores of multiple locations of Alpine halite deposits, flame-sterilized them, dissolved them in sterile saline, and collected bacteria by filtration. Colonies similar to Hcc. salifodinae grew from the samples, but the drill cores yielded fewer colonies than the mines, indicating that drill cores were probably less contaminated. By sequencing the 16S rRNA gene, they found sequences homologous to Hbt. salinarum. Low sequence homology of many DNA clones indicated novel, uncultured taxa. The findings further support the concept of connected Permian-Triassic hypersaline seas in Europe. Using the same flame-sterilization and dissolution techniques, the same group identified another species of haloarchaea, Hcc. dombrowskii (named in honor of Heinz Dombrowski), in a newly blasted tunnel of the Bad Ischl mine [37], and a new species of Halobacterium, Hbt. noricense, from a drill core in a salt mine in Altaussee, Austria [38].
With many of these early studies, it is imperative to note that some of these isolates came from bulk dissolution of rock salt without detailed specified surface sterilization procedures or reported contamination testing. Ethanol is ineffective in killing bacterial endospores. Flaming the surfaces with controlled temperature change on the surface of the halite crystal as described in Radax et al [36] may not achieve sterilization. Therefore, the validity of these results and antiquity of the resulting microorganisms relied more on repeated recovery of similar halophilic taxa across distinct geographic deposits, although contamination by microorganisms common to hypersaline environments cannot be excluded in these early studies.
Refining Extraction Techniques
As the amount of literature exploring the extraction and isolation of halophilic microorganisms progressed throughout the years, researchers refined their techniques to better defend the antiquity of these species. Rosenzweig et al. [17] investigated chemical sterilization techniques and developed a method of reliable surface sterilization by treating halite crystals with 10-M NaOH and 10-M HCl, 5 minutes each. In addition, there was an emphasis on identification and isolation of only primary halite crystals (instead of recrystallized crystals) and aseptic extraction of primary fluid inclusions. Armed with knowledge and skills of surface sterilization, halite crystal selection, and microscopic fluid extraction, the group isolated the famous 2-9-3 clone from the Salado formation, a Permian structure found in New Mexico, USA, and is dated at around 250 million years old by invertebrate fossils and radiometric aging [1]. Strain 2-9-3 is a moderately halophilic bacterium with the 16S rRNA gene 99% homologous to the spore-forming bacterium Virgibacillus marismortui and 97.5% homologous to Virgibacillus pantothenticus. Out of 53 crystals that were obtained from the formation, only 2 contained inclusions from which bacteria were grown, indicating a rare survival occurrence.
Mormile et al. [39] obtained halite samples in a drill core from Death Valley, California, 8 m and 85 m in depth (9,600 and 97,000 years old according to U-Th dating). They examined fluid inclusions with a microscope and found bacterium-like microparticles in the halite inclusions of both depths. Subsequently, they used a microdrill and a micropipette to aseptically extract microparticle-containing fluid directly from primary inclusions of primary halite crystals. A live organism was isolated from 85-m-deep halite but not from 8-m-deep halite. The organism was identified by its 16S rRNA as the archaeon Hbt. salinarum.
Vreeland et al. studied microorganisms in the Sergipe Basin in Brazil, which contained salts from the Aptian stage in the early Cretaceous period dated between 121-112 million years old [40]. Fifty-six primary crystals in total were obtained and tested for fluid inclusion microbial growth after NaOH-HCl treatment, resulting in six microbial isolates from four crystals. Three of the crystals each contained one microbial isolate while one crystal contained three isolates. All six isolates belonged to the Archaea domain, with one from the genus Natronobacterium and five from the genus Halobacterium. Archaeal taxa were determined by lipid analysis along with 16S rRNA sequencing.
Park et al. [41] gathered primary halite crystal samples from three different regions, Remolinos in Spain (Oligocene, 23 million years old), Cretaceous salts of the Sergipe Basin, Brazil (112-121 million years old), and Silurian Salts from the Michigan Basin, USA (416-419 million years old), and analyzed their DNA contents. From these three sites, a total of 103 crystals were sampled. After NaOH-HCl treatment and dissolution of single crystals in extraction buffer, only one crystal from each site was found to contain DNA. PCR with the DNA using 16S rRNA gene-specific primers amplified genes of Halorubrum and Haloarcula from the Oligocene crystal while the Cretaceous and Silurian crystals contained DNA of Halobacterium and unclassified species. These results reinforced previous findings that Halobacterium is the most common haloarchaea in NaOH-HCl-sterilized, more ancient samples. Even though Halococcus species were isolated in earlier European reports, DNA analyses in these studies still pointed to Halobacterium [36]. The authors also described a 55-nucleotide insert in the 16S rRNA in Cretaceous and Silurian samples but not in Oligocene halite. Along with the unclassified taxa amplified from the older samples, the authors argued for different ancient 16S rRNA.
Gramain et al. [42] found that NaOH-HCl treatment failed to completely kill Hcc. saccharolyticus inoculated on the surface of halite crystals. Nor did the procedure eliminate halococcal 16S rRNA on the crystal. Subsequently, they developed a more rigorous method consisting of sequential treatment with 6% sodium hypochlorite, 10 M NaOH, 10.53 M HCl, and 70% ethanol, each for 20 minutes. The method killed Hcc. saccharolyticus on the surface while sparing Hbt. salinarum entombed in inclusions. The group analyzed halophiles in a drill core obtained from the Salar Grande basin of Northern Chile dated to the Pliocene (1.8-5.3 million years old). While the top, recrystallized halite (not surface sterilized due to small sizes) harbored diverse haloarchaea, the authors only isolated Hbt. noricense from surface-sterilized halite crystals deep in the basin and only found Hbt. salinarum-like 16S rRNA sequences in them. They tested endurance of Halobacterium species in halite inclusions and found that Hbt. noricense recovered faster than Hbt. salinarum after 26 months of entombment in nutrient-free inclusions. Halobacterium species survived better than two other common halophiles, Haloquadratum walsbyi and Salinibacter ruber.
The concept that younger halite contains diverse haloarchaea is reinforced by a study of Schubert et al. [43] of a Death Valley salt core. They isolated Halorubrum, Natronomonas, and Haloterrigena from 5 of 881 crystals (surface sterilized with 10 M NaOH) between 10,000 and 34,000 years old. Although the samples were primary crystals 13.0-17.8 m in depth, the microbiological profile was similar to the unsterilized, recrystallized surface halite of the Salar Grande. Using PCR and sequencing, authors from the same laboratory also discovered haloarchaeal 16S rRNA in salt crystals from the same depth of the Death Valley after treating them with 10-M HCl for 20 minutes which successfully destroyed spiked human DNA [44]. The DNA sequences of Halorubrum and Natronomonas were identical to the 16S rRNA extracted from the corresponding isolates. In addition, the crystals also contained sequences related to Haloarcula, Halobiforma, Halosimplex, Halomicrobium, and Halonotius, as well as abundant unclassified haloarchaeal DNA.
In addition to surface-sterilization with concentrated NaOH and HCl, Jaakkola et al. [45] mechanically removed the outer shell of salt crystals by chiseling. They obtained eight haloarchaeal isolates from a rock salt drill core over 600 m in depth in the Jianghan Basin of Hubei Province, China. The sample was determined to be Eocene (38-41 million years old). Based on partial 16S rRNA sequences, two of the isolates were Halobacterium and the other six were Halolamina. Subsequently, the group obtained a 123-million-year-old Cretaceous drill core halite sample (2000-meter in depth) from the same basin [4]. Using the same techniques, they isolated eight haloarchaeal colonies. Intensive genomic and phenotypic analyses demonstrated that the colonies belonged to a Halobacterium species whose 16S rRNA was 99% identical to that of Hbt. noricense, its closest phylogenetic relative. However, based on its low DNA–DNA re-association value (43%) with Hbt. noricense, the authors determined that the Halobacterium was a previously unidentified species and named it Halobacterium hubeinese. The two Halobacterium isolates that were previously obtained from the Eocene sample belonged to the same species (99.9% identity in 16S rRNA).
Meng et al. [46] further studied halophilic organisms in the Jianhan Basin using an Eocene halite sample (33.9-48.6 million years old) of a drill core that was rich in primary inclusions. To ensure surface-sterilization, the authors cracked the crystals and treated them with concentrated NaOH and HCl, 10 minutes each. They dissolved the salt and isolated from it a bacterial species, an Oceanobacillus by 16S rRNA. It was the second member of the spore-forming Bacillaceae family (after the Virgibacillus revived in 2000) to be isolated from ancient halite.
Hoover and Pikuta [47] were the first group to isolate anaerobes (vibrio and rods) from Mississippian (350 Myr) halite as well as sylvite. They UV-irradiated crystals for 10 minutes after treating them in 70% alcohol before extracting the interior by fracturing them aseptically. They found more bacteria in sylvite than in halite.
Direct Microscopy
As early as 1935, Rippel [48] dissolved newly blasted Permian halite crystals in water, stained the residues, and discovered bacterial cells, mostly rods. In 1963, Dombrowski [28] published a high-resolution photomicrograph of a rod-shaped bacterium “embedded in the crystalline structure of the salt” in thin sections. In 2003, Mormile et al. [39] were the first group to photograph bacteria in primary halite inclusions. Schubert et al. [49] provided a detailed morphological study documenting miniaturized cocci less than 1 µm in diameter in primary halite inclusions of 22-Kyr, 25-Kyr, and 34-Kyr halite crystals.
The most recent report on ancient halophiles was published in 2022, documenting microorganisms in the most ancient halite samples ever studied. Using halite crystals from the 830-million-year-old Browne Formation of central Australia, Schreder-Gomes et al. [50] examined bacteria-like structures in situ. A razor blade was used to cleave halite into 1-2-mm-thick sections that were then scrupulously screened to only include primary fluid inclusions. Individual fluid inclusions were then visualized with both transmitted light and ultraviolet-visible light to detect optical appearance and fluorescent response. Through size, shape, and fluorescent response the authors were able to characterize the microorganisms as prokaryotes, eukaryotes, and organic compounds. Although this approach does not establish viability or the age of the observed structures, it provides in situ evidence for microorganism-like structures and organic material within primary fluid inclusions of Precambrian halite. Future improvements in sample preparation and optical technology may permit more detailed characterization of microorganisms or microbial remains in halite crystals.
A chronological list of live microbes isolated from ancient halite is provided in table 1.
Why Geological-Age Survival of Endospores Is Biologically Difficult
The magnitude of the problem is easy to miss when geological ages are treated only as dates. A claim of survival for 100-250 million years requires a cell to preserve or repeatedly repair essential DNA, proteins, membranes, and other structures for intervals many orders of magnitude longer than direct laboratory observations of dormancy. Protection by salt can slow some reactions, but it cannot make spontaneous chemistry, natural radiation, or energy requirements disappear. The following evidence does not establish one universal expiration date for microbial life; instead, it shows why geological-age viability demands an additional mechanism or history of survival than we know currently.
The following sections examine several biological constraints on the survival of microorganisms over geological timescales, beginning with bacterial endospores.
Biological Constraints on Endospore Longevity
An endospore is a dormant form produced mainly by members of Bacillaceae and Clostridia [51]. Its dehydrated core contains the bacterial genome, small acid-soluble proteins, and other molecular components protected by a multilayered coat [52,53]. Extremely long survival times have been proposed for dried bacteria and endospores [29,51]; however, indefinite persistence would require spontaneous molecular damage to remain negligible or repairable over time. Residual water permits slow hydrolytic reactions, while environmental radionuclides can cause direct and indirect damage. There is evidence that dose-rate dependence of ionizing radiation is minimal, meaning lower doses over a longer period can cause similar damage as higher doses over a shorter duration [54]. Temperature accelerates many damaging chemical reactions [55]; for example, dry storage of Bacillus subtilis var. niger spores at 37°C caused substantial loss of viability over weeks [56], whereas higher-temperatures (65-70°C) can damage spores over hours [57]. The thermal history of individual halite deposits therefore matters when evaluating proposed survival intervals [28]. Because dormant spores lack ongoing repair and replacement of biomolecules, accumulated lesions must remain compatible with successful germination and repair during revival [53]. The following sections examine radiation, DNA and protein degradation, and empirical changes in viable-spore abundance in sediments.
Irradiation by 40K Limits Spore Survival Time
In a direct response to the report of strain 2-9-3 from the approximately 250-Myr Salado Formation [1], Kminek et al. examined whether natural ionizing radiation would constrain the persistence of bacterial spores in halite fluid inclusions [58]. Their model treated 40K as the principal radionuclide in the brine inclusion and combined calculated dose accumulation with experimentally measured radiation-inactivation constants for spores, including the halophilic spore-former Virgibacillus marismortui. Assuming a 10-µL inclusion and an initial bacterial concentration of 4000 CFU/mL, the calculated chance of finding a viable spore fell to about 1 in 25,000 after 17–34 Myr at 7-g/L potassium (average concentration in fluid inclusions) and 55–109 Myr at 2.2-g/L potassium (low potassium fluid), depending on the inactivation constant used. The authors therefore concluded that, in the absence of active repair during dormancy, ionizing radiation from 40K within the inclusion body would impose an upper survival limit of less than 109 Myr under the modeled halite conditions.
Degradation of DNA
DNA is a relatively unstable biomolecule and undergoes spontaneous base modification, hydrolysis of phosphodiester and N-glycosidic bonds, oxidation, and alkylation [59]. The N-glycosidic linkage between deoxyribose and purine bases is particularly labile, with depurination strongly dependent on temperature and chemical environment [60,61]. Dormant spores cannot continuously repair such damage; lesions can therefore accumulate until germination, when survival depends on the extent of protection and on successful repair during revival [62]. Johnson et al. [63] examined bacterial persistence in permafrost samples spanning up to approximately one million years by combining long-fragment PCR, uracil-N-glycosylase treatment, and low-temperature respiration measurements. Four-kilobase bacterial DNA was recovered from 400–600-kyr samples but not from samples dated to 740 kyr or approximately 1 Myr. Respiration was also detected in samples younger than 600 kyr but not in the 740-kyr sample. In addition, they found evidence of significant cytosine deamination in low-GC Gram-positive spore-forming bacteria. Notably, the oldest amplifiable DNA was dominated by non-spore-forming Actinobacteria rather than recognized endospore-formers. These observations support bacterial survival to roughly half a million years under the studied permafrost conditions, but they do not establish a universal maximum survival time for bacteria or spores. The authors instead argued that low-level metabolism and DNA repair can be more effective than strict dormancy for very long persistence. This study did not examine halite inclusions, but it illustrates the difficulty of maintaining genome integrity during prolonged dormancy.
Degradation of Proteins
Spontaneous degradation of proteins has also been extensively studied. From measurements of free and peptide-bound amino acids in deep-sea drill cores, Bada and Man [64] estimated a peptide-bond hydrolysis half-life of approximately 1–2 Myr in calcareous sediments, while noting that hydrolysis may be considerably slower in non-carbonate sediments. This is a molecular-diagenesis timescale rather than a direct measurement of microbial viability. Amino-acid racemization provides another cumulative measure of molecular alteration, and aspartate is among the relatively rapidly racemizing residues [65]. An experimental study related aspartate racemization to loss of viability in Geobacillus stearothermophilus spores and extrapolated measured high-temperature kinetics (50-127°C) to environmentally relevant temperatures [12]. For Aarhus Bay conditions of 0–15°C, the model predicted spore half-lives of 40–421 years; for Lake Constance sediment at 4°C, the predicted half-life was about 220 years. These estimates depend on extrapolation and on environmental variables such as temperature, pH, ionic strength, and matrix effects, but they are broadly consistent with viability-based estimates from cold sediments.
Data on high-temperature inactivation of endospores provide an instructive warning about long extrapolations. Nicholson [11] compiled thermal-inactivation data for spores of several Bacillus species and extrapolated them to the estimated 25–40°C temperature range of the Salado Formation. For a bacterial group containing moderately thermophilic B. stearothermophilus strains, the extrapolated D-values ranged from approximately 1.9 × 10⁹ to 1.9 × 10¹² years. Nicholson explicitly noted that the upper values were physically nonsensical and emphasized that no direct thermal-inactivation data were then available for spores in brine. The exercise therefore demonstrates that a mathematically good fit over an experimental temperature range does not guarantee a biologically meaningful extrapolation across geological timescales.
Observed Decline of Spore Counts in Deeper Sediments
Several sediment studies have reported approximately exponential declines in the abundance of culturable or viable endospores with burial age. De Rezende et al. [66] studied thermophilic sulfate-reducing endospores in Aarhus Bay and obtained an estimated culturability half-life of approximately 330 years. A related study of thermophilic fermentative endospores at the same site reported an exponential decrease with depth and an estimated half-life of 350 years [67]. Rothfuss et al. [68] found that viable aerobic and anaerobic heterotrophic spores in Lake Constance sediments decreased with depth and fell below detection in approximately 8,900-year-old sediment. Their reported death-rate constants of 0.0013–0.0025 year⁻¹ correspond to first-order half-lives of approximately 277–533 years. These values describe population-level loss of viability or culturability in sediment and should not be interpreted as universal intrinsic half-lives of individual spores. Wörmer et al. [69] used dipicolinic acid (DPA) as a culture-independent proxy for endospore abundance across a global marine-sediment dataset. A segmented regression showed a breakpoint near 14.2 kyr: endospore concentration declined by more than two orders of magnitude over the younger interval, whereas across the approximately 20 Myr represented by the older segment the DPA-derived concentration was only halved. This latter value is therefore an abundance trend across heterogeneous sediments, not a 20-Myr measured viability half-life. Although DPA is diagnostic of endospores and the authors argued against a major contribution from fossil DPA, DPA quantification itself is not a direct viability assay.
Viable bacterial concentrations in brines have been reported near 4000 CFU/mL [70]. Laboratory experiments indicate that only a minor fraction of haloarchaea survive during incorporation into halite. Kixmüller and Greie [71] observed that approximately 10% of wild-type Hbt. salinarum cells remained viable following entombment in halite, a value they considered consistent with earlier observations by Fendrihan et al. [72] and Kottemann et al. [73]. As an illustrative calculation, suppose that 10% of those cells remain represented as viable spores after entrapment, giving 400 spores/mL of inclusion brine. If fluid inclusions occupy 2% of halite volume [74] and halite density is 2.16 g/cm³, one metric ton of halite contains approximately 9.26 L of inclusion fluid and therefore about 3.7 × 106 spores under these assumptions. Applying the longest viability-associated sediment half-life considered above, 533 years, a simple first-order decay model would reduce the expected number below one after approximately 11,600 years. This calculation is a population-level extrapolation, not a universal survival limit.
The Energy Problem for Live Cells in a Closed Fluid Inclusion
While the lifespan of a dormant endospore is limited by accumulation of unrepaired molecular damage, a cell with any level of metabolism faces a starvation problem when entombed in a small drop of saline. Recent developments in microbiology revealed a couple of survival mechanisms for starved microorganisms, whether they form spores or not.
Long-Term Stationary Phase (LTSP)
The bacterial growth curve consists of five phases: lag phase, log phase, stationary phase, death phase, and long-term stationary phase [7,75]. The long-term stationary phase (LTSP) commences when most of the microbial population dies and releases nutrients into the environment, initiating nutrient recycling by cannibalism [76]. Escherichia coli (E. coli) was found to survive in broth cultures for at least 5 years [75]. During LTSP, microorganisms adapt and gain the ability to survive under various environmental stress. When deeply starved, most Bacillus subtilis cells sporulate, but others assume a slow-growing coccoid shape with their doubling time prolonged from ~40 minutes to ~4 days [77]. The DNA-binding Protein from Starved cells (Dps) becomes the most represented protein in E. coli cells and protects the cell and its DNA from ensuing oxidative stress [78]. Homologues of Dps have been found in other bacteria and archaea [79,80,81].
To examine persistence of living cells under extreme energy limitation, Shoemaker et al. [82] followed 21 heterotrophic bacterial taxa in long-term stationary phase at 25°C in darkness and modeled mortality trajectories. The resulting estimates of population time to extinction spanned roughly 1–100,000 years, with large differences among taxa and especially long persistence among spore-forming Bacillus. These are model-based population-extinction times from laboratory starvation experiments rather than direct measurements of continuous survival over those intervals. Gray et al. [77] further showed that long-term Bacillus subtilis populations can persist without repeated wholesale cycles of sporulation and germination, underscoring that LTSP survival is a population-dynamic process distinct from simple endospore dormancy.
It is important to note the thermodynamic cost of cannibalism in LTSP. Schink et al. [76] found that it took 5.5 dead E. coli cells to make one new living cell. When E. coli enters LTSP, numerous mutations that increase the efficiency of energy use (growth advantage in stationary phase, GASP) are selected, most notably those inactivating the rpoS gene that codes the general stress response sigma factor. By partially inactivating rpoS, these mutants demonstrate enhanced catabolism of amino acids as alternative carbon sources. Paradoxically, ΔrpoS mutant cells increase the death rate by 1.8-fold and increase the number of dead cells to build a new cell to 10, essentially creating a bacterial population that extinguishes quicker [76,83]. Because of the improvisational nature of natural selection, GASP may only mean transient advantage instead of long-term survival. Indeed, carbon-efficient E. coli cells out of LTSP demonstrated reduced survival time in the presence of a producer organism, Chlamydomonas reinhardtii [84].
Assuming a bacterial concentration of 4000 CFU/mL [70], the number of organisms originally trapped in primary inclusions should be small. If we assume a generous number of 100 organisms in a fluid inclusion and it takes 5 dead cells to grow a live cell, it only takes three cycles to reduce the number of live cells to less than Thus, cannibalistic recycling alone appears insufficient to sustain a small, closed halophile population for millions of years in a brine inclusion.
Haloarchaeal Microspheres
Although archaea do not form endospores, they are known to endure long-term starvation which may have to do with nutrients stored in their polyploid genome [45]. Haloarchaea were revived from Dead Sea brine samples enriched with 1% peptone after 56 years of laboratory storage [85]. Like many microbes and plants [86,87], archaeal cells undergo extensive morphological and biochemical changes under severe starvation. Schubert et al. [49] reported miniaturized archaea in Death Valley salt cores. In the laboratory, Fendrihan et al. [88] observed the formation of 0.4-µm spherical particles when rod-shaped Halobacterium species were placed into a setting of reduced external water activity. Unlike endospores, haloarchaea spheres can form quickly, three to four from each rod, presumably due to their polyploid genome. These spherical Halobacterium remained viable in laboratory conditions for at least 6 years without organic nutrients and can reverse to rod shape when water activity increases. While the rod and spherical forms of the Halobacterium genus displayed many of the same biochemical properties, there was a markedly large difference in ATP content, with the microspheres containing approximately 50-fold less ATP than the rods. These observations are consistent with a strongly reduced, dormancy-like physiological state. Schubert et al. [43] revived haloarchaea microspheres from ancient Death Valley samples. Compared to the 240-fold reduction of ATP content in bacterial endospores [89], haloarchaeal microspheres may retain some metabolic activity, although ATP content alone does not establish the rate or physiological purpose of that activity. Besides their polyploid DNA, haloarchaeal microspheres may not possess more mechanisms of survival than non-spore-forming bacteria in LTSP. Pseudomonas aeruginosa, for example, was found to survive for long periods of time (at least 14 years according to Elabed et al.) in the inclusions of lab-grown halite crystals [90,91], in partial vindication of Heinz Dombrowski.
Haloarchaeal microspheres can coexist with cells or remains of the alga Dunaliella in brine inclusions [92,93]. Dunaliella can accumulate very high intracellular glycerol concentrations, and glycerol has been proposed as an important carbon and energy source for co-trapped haloarchaea [6]. A published energy-budget calculation based on a 10-µm Dunaliella cell containing 5.5 M glycerol and a miniaturized 10-fg-C prokaryotic cell estimated that this carbon reserve could support repair metabolism for approximately 12 Myr under a maintenance metabolic rate of 10−7 hr−1 [92]. The assumed rate was derived from the community-level metabolic rates compiled by Price and Sowers [94], in which rates on the order of 10-7 h-1 fall within the range attributed to maintenance metabolism at subfreezing temperatures. This corresponds to only 9.1 x 10-21 W per cell. Subsequently, Kempes et al. [95] adopted 3.17×10-20 W cell⁻¹ as a lower-bound estimate of the basal power requirement, based on the minimum cell-specific energy turnover inferred by Lever et al. [87] for sulfate-reducing communities in energy-limited subsurface sediments. Other directly inferred maintenance energy requirements were orders of magnitude higher [96,97]. Thus, the 12-Myr estimate represents an assumption-sensitive upper bound of survival time in a small inclusion body with a coexisting algal cell. In addition, the usable energy yield depends on redox conditions and metabolic pathways within the inclusion. Anoxia and acid accumulation via fermentation could shorten the time further.
Table 2 summarizes quantitative estimates relevant to microbial survival time over geological timescales.
The Molecular-Age Paradox: Why Do Ancient Isolates Look Modern?
Molecular-clock approaches, which estimate divergence from sequence change calibrated against time, can be informative for studying the evolution of microorganisms for which fossil records yield limited information [98]. Early calibrations of 16S rRNA sequence divergence estimated rates at about 1% divergence in the span of 50 million years using diverse evolutionary timeframes such as cyanobacterial fossils and the divergence of mitochondria and chloroplasts from related prokaryotes [99]. A more recent calibration of the 16S rRNA clock using fossils of endosymbiotic insect hosts found a similar rate of 1–2% divergence per 50 Ma [100]. Because of the conservation of ribosomal structures, this rate was found to be comparable across prokaryotes and eukaryotes [99].
Even though archaea reproduce slower than bacteria under optimum laboratory conditions, their generation times in the wild are primarily determined by environmental factors such as nutrient availability, temperature, and bacteriophages [101,102,103]. Most bacteria in the wild are in long-term stationary phase [104,105,106]. Based on absorption of 3H-labelled thymidine, the growth rates of halophilic archaea were found to be comparable to those of bacteria in saltern ponds of various salinity levels [107,108]. In the higher temperatures of hypersaline environments, halophilic archaea can achieve faster growth than many bacteria, with doubling times between 1.5 and 3 hours [102].
On the other hand, observed per-generation genome-wide mutation rates of halophilic archaea are also comparable to those of mesophilic bacteria [109,110]. Even though polyploidy of archaea may facilitate recombinational DNA repair, polyploidy also relaxes natural selection and has been found to facilitate accumulation of deleterious mutations in plants [111]. Genomic streamlining favors deletion in multicopy genes [112,113]. Halobacterium salinarum and Haloferax volcanii both feature high rates of indel mutations with a bias toward deletions [110,114]. These observations do not provide a clear empirical basis for assuming that haloarchaea as a group possess exceptionally slow molecular-evolution rates.
Nevertheless, DNA sequences of ancient archaeal isolates are surprisingly similar to their modern relatives. For example, McGenity et al. [115] noted that 16S rRNA similarity between an isolate from Upper Devonian halite (~240 million years old) and the modern archaeon Hbt. saccharovorum was difficult to reconcile with the geological age assigned to the host halite. Likewise, the 250-million-year-old Hcc. salifodinae is 98.9% identical in 16S rRNA to modern Hcc. saccharolyticus [98]. Their cell wall morphology and biochemistry are also strikingly similar.
Members of the Haloarcula genus have two or three paralogous copies of 16S rRNA in their genomes. Because paralogous rRNA genes can preserve information about their divergence history, comparisons among copies provide an additional perspective on the age of genes and organisms. The reported similarity among paralogs in the putatively ancient material fell within the range observed in modern archaeal genomes [116].
If the recovered lineages had remained genetically isolated for very long intervals, one might expect distinctive phylogenetic patterns. Instead, ancient archaeal isolates are scattered among major modern genera and are often closer to modern organisms than to each other [117]. Likewise, modern archaeal sequences do not consistently differ more from the putatively ancient sequences than they do from one another [116].
On the bacterial side, 16S rRNA sequences amplified from Upper Silurian halite (415-425 Myr) were 98.4-99% homologous to a modern β-proteobacterium, Aquabacterium commune [118]. Likewise, the Bacillus strain isolated from 250-Myr Permian salt, strain 2-9-3, was 99% identical in 16S rRNA to modern Virgibacillus marismortui from the Dead Sea [1]. To further compare the DNA of 2-9-3 and Virgibacillus marismortui, Maughan et al. [119] sequenced two protein-coding genes, recA and splB, in both strains. The recA gene is conserved throughout the bacterial domain, while the latter is unique to Gram-positive spore-formers. The two organisms demonstrated one synonymous substitution among the 404 nucleotides of the recA gene and two synonymous mutations among the 619 nucleotides of the splB gene. In other words, they share identical protein sequences. The RecA and SplB proteins of other Bacillus species are different from those of 2-9-3 to various degrees. The extremely small number of differences in these two protein-coding genes therefore remains difficult to reconcile with prolonged genetic isolation over approximately 250 million years. A simple linear extrapolation across more divergent Bacillus species would not be justified because substitution rates vary among genes and lineages.
Biomedical Significance
Although the organisms discussed here are environmental microbes rather than human cells, the underlying constraints are familiar to biomedical science: spontaneous DNA lesions, protein damage, oxidative and radiation injury, energy-dependent repair, stress responses, and the tradeoff between dormancy and maintenance. Ancient halite pushes these processes toward an extreme limit. If some isolates truly persisted for geological intervals, their biology would imply unusually effective strategies for preserving molecular integrity at vanishingly low energy flux. If they are younger than the host deposits, determining how they entered apparently primary inclusions becomes an equally important problem in contamination control and interpretation of ancient biological material.
Possible Explanations
Contamination by Modern Organisms
Early studies were more vulnerable to contamination than later investigations because many relied on bulk brine, efflorescence, or whole-crystal dissolution and used surface treatments that would not meet current standards. Some early isolates may therefore have represented modern halophiles introduced during sampling or processing. Ethanol immersion and brief flaming, although useful decontamination measures, may not reliably eliminate all surface microorganisms. Contamination remains a plausible explanation for some reports, but it becomes less satisfactory for studies using deeply buried material, rigorous surface sterilization, petrographic screening, and direct extraction of primary fluid inclusions.
Recrystallization and Groundwater Infiltration
Halite formations can undergo ductile deformation under pressure, accompanied by solution-precipitation, folding, diapirism, and recrystallization [74,120,121]. During such alteration, fluid inclusions may migrate, leak, or be redistributed, and new inclusions can form during growth of recrystallized halite. Such inclusions may be primary with respect to the newly grown crystal without being syn-depositional with the original salt bed. Stable-isotope studies provided evidence consistent with mixing of evaporite brines and meteoric groundwater in some salt deposits [120,122]. The contribution of meteoric water generally decreases with depth in the studied systems. Selecting deeply buried crystals that retain primary growth features, such as chevrons, and then targeting demonstrably primary inclusions greatly reduces this concern and has become an important control in paleomicrobiological studies. Nevertheless, petrographic identification of primary growth features cannot by itself establish that microorganisms presently enclosed within an inclusion have remained isolated since the original depositional event. Recrystallization and groundwater infiltration therefore remain possible explanations in particular deposits but are less satisfactory where primary depositional textures and inclusion chemistry are well preserved.
Unknown Survival Mechanisms
Although bacterial endospores have been extensively studied, much less is known about long-term stationary-phase survival and the dormancy-like microspheres formed by haloarchaea. Extremophiles repeatedly reveal physiological adaptations that expand known limits of life, so additional survival mechanisms cannot be excluded. One possibility is that cells enter extremely low-activity survival states punctuated by rare episodes of repair or maintenance before accumulated molecular damage becomes irreversible. Such episodic metabolism could, in principle, reduce the energetic cost relative to continuous maintenance while permitting molecular repair, but direct evidence for the existence of such a strategy and its ability to sustain viability over geological timescales is lacking.
Genomic Divergence Underestimated by 16S rRNA
Most molecular comparisons between putatively ancient and modern microorganisms have emphasized 16S rRNA genes because they are highly conserved. DNA-DNA hybridization provides evidence that this marker can understate broader genomic differentiation. Legat et al. [98] reported only 63.6% DNA-DNA relatedness between the Permian isolate Hcc. salifodinae BIpT and Hcc. saccharolyticus DSM 5350T despite 98.9% 16S rRNA gene identity. This hybridization value is a measure of genomic relatedness rather than direct nucleotide-sequence identity. Park et al. [41] reported a 55-nt insertion in 16S rRNA gene sequences recovered from Cretaceous and Silurian halite that was absent from their Oligocene sample, and therefore not included in aligned comparisons. These observations caution against equating high 16S rRNA similarity with genome-wide identity. As additional ancient isolates are subjected to whole-genome sequencing, the extent of genomic divergence from modern relatives may prove greater than is apparent from 16S rRNA comparisons alone.
Conclusions
The evidence reviewed here leaves a genuine two-sided problem. Modern contamination is a credible explanation for some early reports, but it becomes less persuasive for studies using deeply buried material, rigorous surface sterilization, petrographic selection of primary crystals, and direct sampling of fluid inclusions. At the same time, no established mechanism readily explains preservation of viable cells for tens to hundreds of millions of years in a small, nutrient-limited inclusion while spontaneous molecular damage continues to accumulate.
Resolving the paradox requires experiments that constrain both geological history and biological history in the same specimen. Future studies should combine documented primary halite and inclusion petrography with modern contamination controls, direct in situ imaging and chemistry, cultivation where possible, and whole-genome analysis of recovered organisms. For biomedical science, the broader significance is the boundary condition these claims place on cellular longevity: a viable cell must retain enough molecular information and repair capacity to restart life after prolonged metabolic restriction. Whether ancient halite ultimately reveals extraordinary persistence or an unrecognized route of later entry, it remains a stringent natural test of our understanding of dormancy, molecular maintenance, and the limits of cellular survival.

Author Contributions

Yingguang Liu initiated the writing project. Colby Renshaw conducted initial literature search and provided the initial draft. Yingguang Liu did most of the revisions and the mathematical analyses.

Funding

Publication of this review is funded by Liberty University College of Osteopathic Medicine.

References

  1. Vreeland, R.H.; Rosenzweig, W.D.; Powers, D.W. Isolation of a 250 million-year-old halotolerant bacterium from a primary salt crystal. Nature 2000, 407, 897–900. [Google Scholar] [CrossRef]
  2. Nickle, D.C.; Learn, G.H.; Rain, M.W.; Mullins, J.I.; Mittler, J.E. Curiously modern DNA for a ‘250 million-year-old’ bacterium. J. Mol. Evol. 2002, 54, 134–137. [Google Scholar] [CrossRef]
  3. Becker, E.A.; Seitzer, P.M.; Tritt, A.; et al. Phylogenetically driven sequencing of extremely halophilic archaea reveals strategies for static and dynamic osmo-response. PLoS Genet. 2014, 10, e1004784. [Google Scholar] [CrossRef]
  4. Jaakkola, S.T.; Pfeiffer, F.; Ravantti, J.J.; et al. The complete genome of a viable archaeum isolated from 123-million-year-old rock salt. Environ. Microbiol. 2016, 18, 565–579. [Google Scholar] [CrossRef]
  5. Soto, L.; DasSarma, P.; Anton, B.P.; et al. Genome sequence of an extremely halophilic archaeon isolated from Permian Period halite, Salado Formation in New Mexico, USA: Halobacterium sp. strain NMX12-Microbiol Resour. Announc. 2024, 13, e00778-24. [Google Scholar] [CrossRef]
  6. Lowenstein, T.K.; Schubert, B.A.; Timofeeff, M.N. Microbial communities in fluid inclusions and long-term survival in halite. GSA Today 2011, 21, 4–9. [Google Scholar] [CrossRef]
  7. Pletnev, P.; Osterman, I.; Sergiev, P.; Bogdanov, A.; Dontsova, O. Survival guide: Escherichia coli in the stationary phase. Acta Naturae 2015, 7, 22–33. Available online: https://pubmed.ncbi.nlm.nih.gov/26798489/. [CrossRef]
  8. Stan-Lotter, H.; Fendrihan, S. Halophilic archaea: life with desiccation, radiation and oligotrophy over geological times. Life 2015, 5, 1487–1496. [Google Scholar] [CrossRef]
  9. Vuillemin, A.; Wankel, S.D.; Coskun, Ö.K.; et al. Archaea dominate oxic subseafloor communities over multimillion-year time scales. Sci. Adv. 2019, 5, eaaw4108. [Google Scholar] [CrossRef]
  10. Bradley, J.A.; Arndt, S.; Amend, J.P.; Burwicz, E.; Dale, A.W.; Egger, M.; LaRowe, D.E. Widespread energy limitation to life in global subseafloor sediments. Sci. Adv. 2020, 6, eaba0697. [Google Scholar] [CrossRef]
  11. Nicholson, W.L. Using thermal inactivation kinetics to calculate the probability of extreme spore longevity: implications for paleomicrobiology and lithopanspermia. Orig. Life Evol. Biosph. 2003, 33, 621–631. [Google Scholar] [CrossRef]
  12. Liang, R.; Lau, M.C.; Baars, O.; Robb, F.T.; Onstott, T.C. Aspartic acid racemization constrains long-term viability and longevity of endospores. FEMS Microbiol. Ecol. 2019, 95, fiz132. [Google Scholar] [CrossRef]
  13. Hunsche, U.; Hampel, A. Rock salt—the mechanical properties of the host rock material for a radioactive waste repository. Eng. Geol. 1999, 52, 271–291. [Google Scholar] [CrossRef]
  14. Martinez-Donate, A.; Moscardelli, L.; Ko, L.; Melani, L.; Schuba, N.; Bhattacharya, S.; Ruiz Maraggi, L. Geological and geochemical characterization of salt-bearing sequences for hydrogen storage in the Delaware Basin (West Texas). 4th EAGE Global Energy Transition Conference and Exhibition, GET 2023, 2023; 2023, pp. 1–5. [Google Scholar] [CrossRef]
  15. Ruiz Maraggi, L.M.; Martinez Doñate, A.; Melani, L.; Moscardelli, L.G. Hydrogen storage potential of Salado Formation in the Permian Basin of West Texas, United States. Int. J. Hydrogen Energy 2025, 144, 496–512. [Google Scholar] [CrossRef]
  16. Chen, J.; Ren, S.; Yang, C.; Jiang, D.; Li, L. Self-healing characteristics of damaged rock salt under different healing conditions. Materials 2013, 6, 3438–3450. [Google Scholar] [CrossRef]
  17. Rosenzweig, W.D.; Peterson, J.; Woish, J.; Vreeland, R.H. Development of a protocol to retrieve microorganisms from ancient salt crystals. Geomicrobiol. J. 2000, 17, 185–192. [Google Scholar] [CrossRef]
  18. Norton, C.F.; Grant, W.D. Survival of halobacteria within fluid inclusions in salt crystals. Microbiology 1988, 134, 1365–1373. [Google Scholar] [CrossRef]
  19. Rankin, A.H. Fluid inclusions. In Encyclopedia of Geology; Elsevier, 2005; pp. 253–260. [Google Scholar]
  20. Spencer, R.J.; Hardie, L.A. Control of seawater composition by mixing of river waters and mid-ocean ridge hydrothermal brines. In Fluid-Mineral Interactions: A Tribute to H. P. Eugster.; Spencer, R.J., Chou, I.-M., Eds.; The Geochemical Society, 1990; pp. 409–419. Available online: https://www.geochemsoc.org/files/2914/1261/1780/SP-2_409-420_Spencer.pdf.
  21. Hardie, L.A. Secular variation in seawater chemistry: an explanation for the coupled secular variation in the mineralogies of marine limestones and potash evaporites over the past 600 m.y. Geology 1996, 24, 279–283. [Google Scholar] [CrossRef]
  22. Timofeeff, M.N.; Lowenstein, T.K.; Silva, M.A.M.; Harris, N.B. Secular variation in the major-ion chemistry of seawater: evidence from fluid inclusions in Cretaceous halites. Geochim Cosmochim. Acta 2006, 70, 1977–1994. [Google Scholar] [CrossRef]
  23. Johnson, W.J.; Goldstein, R.H. Cambrian sea water preserved as inclusions in marine low-magnesium calcite cement. Nature 1993, 362, 335–337. [Google Scholar] [CrossRef]
  24. Benison, K.C.; Goldstein, R.H. Permian paleoclimate data from fluid inclusions in halite. Chem. Geol. 1999, 154, 113–132. [Google Scholar] [CrossRef]
  25. Bekaert, D.; Avice, G.; Marty, B. Fluid inclusions: tiny windows into global paleo-environments. Commun. Earth Environ. 2025, 6, 820. [Google Scholar] [CrossRef]
  26. Reiser, R.; Tasch, P. Investigation of the viability of osmophile bacteria of great geological age. Trans. Kans. Acad. Sci. 1960, 63, 31–34. [Google Scholar] [CrossRef]
  27. Lowenstein, T.K.; Timofeeff, M.N.; Kovalevych, V.M.; Horita, J. The major-ion composition of Permian seawater. Geochim Cosmochim. Acta 2005, 69, 1701–1719. [Google Scholar] [CrossRef]
  28. Dombrowski, H. Bacteria from paleozoic salt deposits. Ann. N Y Acad. Sci. 1963, 108, 453–460. [Google Scholar] [CrossRef]
  29. Dombrowski, H. Über biogene Formelemente in Mineralwässern und ihre Herkunft. In Wissenschaftliche Arbeiten aus dem Burgenland; 1965; vol 30, pp. 15–28. [Google Scholar]
  30. De Ley, J.; Kersters, K.; Park, I.W. Molecular-biological and taxonomic studies on Pseudomonas halocrenaea, a bacterium from Permian salt deposits. Antonie Van. Leeuwenhoek 1966, 32, 315–331. [Google Scholar] [CrossRef]
  31. Bibo, F.J.; Söngen, R.; Fresenius, R.E. Vermehrungsfähige Mikroorganismen in Steinsalz aus primären Lagerstätten. Kali Steinsalz 1983, 8, 367–373. [Google Scholar]
  32. Norton, C.F.; McGenity, T.J.; Grant, W.D. Archaeal halophiles (halobacteria) from two British salt mines. J. Gen. Microbiol. 1993, 139, 1077–1081. [Google Scholar] [CrossRef]
  33. Denner, E.B.; McGenity, T.J.; Busse, H.J.; Grant, W.D.; Wanner, G.; Stan-Lotter, H. Halococcus salifodinae sp. nov., an archaeal isolate from an Austrian salt mine. Int. J. Syst. Bacteriol. 1994, 44, 774–780. [Google Scholar] [CrossRef]
  34. Stan-Lotter, H.; Sulzner, M.; Egelseer, E.; Norton, C.F.; Hochstein. LI: Comparison of membrane ATPases from extreme halophiles isolated from ancient salt deposits. Orig. Life Evol. Biosph. 1993, 23, 53–64. [Google Scholar] [CrossRef]
  35. Stan-Lotter, H.; McGenity, T.J.; Legat, A.; Denner, E.B.; Glaser, K.; Stetter, K.O.; Wanner, G. Very similar strains of Halococcus salifodinae are found in geographically separated Permo-Triassic salt deposits. Microbiology 1999, 145, 3565–3574. [Google Scholar] [CrossRef]
  36. Radax, C.; Gruber, C.; Stan-Lotter, H. Novel haloarchaeal 16S rRNA gene sequences from Alpine Permo-Triassic rock salt. Extremophiles 2001, 5, 221–228. [Google Scholar] [CrossRef]
  37. Stan-Lotter, H.; Pfaffenhuemer, M.; Legat, A.; Busse, H.J.; Radax, C.; Gruber, C. Halococcus dombrowskii sp. nov., an archaeal isolate from a Permian alpine salt deposit. Int. J. Syst. Evol. Microbiol. 2002, 52, 1807–1814. [Google Scholar] [CrossRef]
  38. Gruber, C.; Legat, A.; Pfaffenhuemer, M.; Radax, C.; Weidler, G.; Busse, H.J.; Stan-Lotter, H. Halobacterium noricense sp. nov., an archaeal isolate from a bore core of an alpine Permian salt deposit, classification of Halobacterium sp. NRC-1 as a strain of H. salinarum and emended description of H. salinarum. Extremophiles 2004, 8, 431–439. [Google Scholar] [CrossRef]
  39. Mormile, M.R.; Biesen, M.A.; Gutierrez, M.C.; Ventosa, A.; Pavlovich, J.B.; Onstott, T.C.; Fredrickson, J.K. Isolation of Halobacterium salinarum retrieved directly from halite brine inclusions. Environ. Microbiol. 2003, 5, 1094–1102. [Google Scholar] [CrossRef]
  40. Vreeland, R.H.; Jones, J.; Monson, A.; et al. Isolation of live Cretaceous (121–112 million years old) halophilic archaea from primary salt crystals. Geomicrobiol. J. 2007, 24, 275–282. [Google Scholar] [CrossRef]
  41. Park, J.S.; Vreeland, R.H.; Cho, B.C.; Lowenstein, T.K.; Timofeeff, M.N.; Rosenzweig, W.D. Haloarchaeal diversity in 23, 121 and 419 MYA salts. Geobiology 2009, 7, 515–523. [Google Scholar] [CrossRef]
  42. Gramain, A.; Díaz, G.C.; Demergasso, C.; Lowenstein, T.K.; Mcgenity, T.J. Archaeal diversity along a subterranean salt core from the Salar Grande (Chile). Environ. Microbiol. 2011, 13, 2105–2121. [Google Scholar] [CrossRef]
  43. Schubert, B.A.; Lowenstein, T.K.; Timofeeff, M.N.; Parker, M.A. Halophilic archaea cultured from ancient halite, Death Valley, California. Environ. Microbiol. 2010, 12, 440–454. [Google Scholar] [CrossRef]
  44. Sankaranarayanan, K.; Lowenstein, T.K.; Timofeeff, M.N.; Schubert, B.A.; Lum, J.K. Characterization of ancient DNA supports long-term survival of haloarchaea. Astrobiology 2014, 14, 553–560. [Google Scholar] [CrossRef]
  45. Jaakkola, S.T.; Zerulla, K.; Guo, Q.; et al. Halophilic archaea cultivated from surface-sterilized middle-late Eocene rock salt are polyploid. PLoS ONE 2014, 9, e110533. [Google Scholar] [CrossRef]
  46. Meng, F.W.; Wang, X.Q.; Ni, P.; et al. A newly isolated haloalkaliphilic bacterium from middle-late Eocene halite formed in salt lakes in China. Carbonates Evaporites 2015, 30, 321–330. [Google Scholar] [CrossRef]
  47. Hoover, R.B.; Pikuta, E.V. Microscopic and microbiological investigations of Mississippian sylvite. Proc. SPIE 2012, 8521, 852108. [Google Scholar] [CrossRef]
  48. Rippel A: Fossile Mikroorganismen in einem permischen Salzlager. Arch. Mikrobiol. 1935, 6, 350–358. [CrossRef]
  49. Schubert, B.A.; Lowenstein, T.K.; Timofeeff, M.N. Microscopic identification of prokaryotes in modern and ancient halite, Saline Valley and Death Valley, California. Astrobiology 2009, 9, 467–482. [Google Scholar] [CrossRef]
  50. Schreder-Gomes, S.I.; Benison, K.C.; Bernau, J.A. 830-million-year-old microorganisms in primary fluid inclusions in halite. Geology 2022, 50, 918–922. [Google Scholar] [CrossRef]
  51. Berman, J.J. Class Bacilli plus class Clostridia. In Taxonomic Guide to Infectious Diseases; Berman, J.J., Ed.; Academic Press, 2012; pp. 65–71. [Google Scholar] [CrossRef]
  52. Mckenney, P.T.; Driks, A.; Eichenberger, P. The Bacillus subtilis endospore: assembly and functions of the multilayered coat. Nat. Rev. Microbiol. 2013, 11, 33–44. [Google Scholar] [CrossRef]
  53. Moir, A.; Cooper, G. Spore germination. Microbiol. Spectr. 2015, 3. [Google Scholar] [CrossRef]
  54. Sivinski, H.D.; Reynolds, M.C. Synergistic characteristics of thermoradiation sterilization. In Life Sciences and Space Research, vol X: Proceedings of the Open Meeting of Working Group 5 of the Fourteenth Plenary Meeting of COSPAR; Akademie-Verlag, 1972; pp. 33–44. Available online: https://ntrs.nasa.gov/api/citations/19870003733/downloads/19870003733.pdf.
  55. Zhang, P.; Setlow, P.; Li, Y. Characterization of single heat-activated Bacillus spores using laser tweezers Raman spectroscopy. Opt. Express 2009, 17, 16480–16491. [Google Scholar] [CrossRef]
  56. Molin G: Inactivation of bacillus spores in dry systems at low and high temperatures. J. Gen. Microbiol. 1977, 101, 227–231. [CrossRef]
  57. Wen, J.; Smelt, J.P.; Vischer, N.O.; Vos, A.L.; Setlow, P.; Brul, S. Heat activation and inactivation of bacterial spores: is there an overlap? Appl. Environ. Microbiol. 2022, 88, e02324-21. [Google Scholar] [CrossRef]
  58. Kminek, G.; Bada, J.L.; Pogliano, K.; Ward, J.F. Radiation-dependent limit for the viability of bacterial spores in halite fluid inclusions and on Mars. Radiat. Res. 2003, 159, 722–729. [Google Scholar] [CrossRef]
  59. Lindahl, T. Instability and decay of the primary structure of DNA. Nature 1993, 362, 709–715. [Google Scholar] [CrossRef]
  60. Gates, K.S. An overview of chemical processes that damage cellular DNA: spontaneous hydrolysis, alkylation, and reactions with radicals. Chem. Res. Toxicol. 2009, 22, 1747–1760. [Google Scholar] [CrossRef]
  61. Allentoft, M.E.; Collins, M.; Harker, D.; et al. The half-life of DNA in bone: measuring decay kinetics in 158 dated fossils. Proc. Biol. Sci. 2012, 279, 4724–4733. [Google Scholar] [CrossRef]
  62. Nicholson, W.L.; Munakata, N.; Horneck, G.; Melosh, H.J.; Setlow, P. Resistance of Bacillus endospores to extreme terrestrial and extraterrestrial environments. Microbiol. Mol. Biol. Rev. 2000, 64, 548–572. [Google Scholar] [CrossRef]
  63. Johnson, S.S.; Hebsgaard, M.B.; Christensen, T.R.; et al. Ancient bacteria show evidence of DNA repair. Proc. Natl. Acad. Sci. U S A 2007, 104, 14401–14405. [Google Scholar] [CrossRef]
  64. Bada, J.L.; Man, E.H. Amino acid diagenesis in deep sea drilling project cores: Kinetics and mechanisms of some reactions and their applications in geochronology and in paleotemperature and heat flow determinations. Earth Sci. Rev. 1980, 16, 21–55. [Google Scholar] [CrossRef]
  65. Demarchi, B.; Collins, M.; Bergström, E.; Dowle, A.; Penkman, K.; Thomas-Oates, J.; Wilson, J. New experimental evidence for in-chain amino acid racemization of serine in a model peptide. Quat. Geochronol. 2013, 16, 158–172. [Google Scholar] [CrossRef]
  66. Rezende, J.R.; Kjeldsen, K.U.; Hubert, C.R.; Finster, K.; Loy, A.; Jørgensen, B.B. Dispersal of thermophilic Desulfotomaculum endospores into Baltic Sea sediments over thousands of years. ISME J. 2013, 7, 72–84. [Google Scholar] [CrossRef]
  67. Volpi, M.; Lomstein, B.A.; Sichert, A.; et al. Thermophilic endospores in cold marine sediments: spore influx from warm geological settings and their contribution to the microbial rare biosphere. Front Microbiol. 2017, 8, 131. [Google Scholar] [CrossRef]
  68. Rothfuss, F.; Bender, M.; Conrad, R. Survival and activity of bacteria in a deep, aged lake sediment (Lake Constance). Microb. Ecol. 1997, 33, 69–77. [Google Scholar] [CrossRef]
  69. Wörmer, L.; Hoshino, T.; Bowles, M.W.; et al. Microbial dormancy in the marine subsurface: global endospore abundance and response to burial. Sci. Adv. 2019, 5, eaav1024. [Google Scholar] [CrossRef]
  70. Vreeland, R.H.; Piselli, A.F.; McDonnough, S.; Meyers, S.S. Distribution and diversity of halophilic bacteria in a subsurface salt formation. Extremophiles 1998, 2, 321–331. [Google Scholar] [CrossRef]
  71. Kixmüller, D.; Greie, J.C. An ATP-driven potassium pump promotes long-term survival of Halobacterium salinarum within salt crystals. Environ. Microbiol. Rep. 2012, 4, 234–241. [Google Scholar] [CrossRef]
  72. Fendrihan, S.; Leuko, S.; Stan-Lotter, H. Effects of embedding Halobacterium sp. NRC-1 in salt crystals and potential implications for long-term preservation. In Proceedings of the III European Workshop on Exo-Astrobiology: Mars: The Search for Life; Harris, R.A., Ouwehand, L., Eds.; European Space Agency, 2004; pp. 203–204. [Google Scholar]
  73. Kottemann, M.; Kish, A.; Iloanusi, C.; Bjork, S.; DiRuggiero, J. Physiological responses of the halophilic archaeon Halobacterium sp. strain NRC1 to desiccation and gamma irradiation. Extremophiles 2005, 9, 219–227. [Google Scholar] [CrossRef]
  74. Roedder E: The fluids in salt. Am. Mineral. 1984, 69, 413–439. Available online: https://pubs.usgs.gov/publication/70014044.
  75. Finkel, S.E. Long-term survival during stationary phase: evolution and the GASP phenotype. Nat. Rev. Microbiol. 2006, 4, 113–120. [Google Scholar] [CrossRef]
  76. Schink, S.J.; Biselli, E.; Ammar, C.; Gerland, U. Death rate of E. coli during starvation is set by maintenance cost and biomass recycling. Cell Syst. 2019, 9, 64–73.e3. [Google Scholar] [CrossRef]
  77. Gray, D.A.; Dugar, G.; Gamba, P.; Strahl, H.; Jonker, M.J.; Hamoen, L.W. Extreme slow growth as alternative strategy to survive deep starvation in bacteria. Nat. Commun. 2019, 10, 890. [Google Scholar] [CrossRef]
  78. Almirón, M.; Link, A.J.; Furlong, D.; Kolter, R. A novel DNA-binding protein with regulatory and protective roles in starved Escherichia coli. Genes Dev. 1992, 6, 2646–2654. [Google Scholar] [CrossRef]
  79. Antelmann, H.; Engelmann, S.; Schmid, R.; Sorokin, A.; Lapidus, A.; Hecker, M. Expression of a stress- and starvation-induced dps/pexB-homologous gene is controlled by the alternative sigma factor sigmaB in Bacillus subtilis. J. Bacteriol. 1997, 179, 7251–7256. [Google Scholar] [CrossRef]
  80. Reindel, S.; Anemüller, S.; Sawaryn, A.; Matzanke, B.F. The dpsa-homologue of the archaeon halobacterium salinarum is a ferritin. Biochim Biophys. Acta 2002, 1598, 140–146. [Google Scholar] [CrossRef]
  81. Wiedenheft, B.; Mosolf, J.; Willits, D.; Yeager, M.; Dryden, K.A.; Young, M.; Douglas, T. An archaeal antioxidant: characterization of a Dps-like protein from Sulfolobus solfataricus. Proc. Natl. Acad. Sci. U S A 2005, 102, 10551–10556. [Google Scholar] [CrossRef]
  82. Shoemaker, W.R.; Jones, S.E.; Muscarella, M.E.; Behringer, M.G.; Lehmkuhl, B.K.; Lennon, J.T. Microbial population dynamics and evolutionary outcomes under extreme energy limitation. Proc. Natl. Acad. Sci. U S A 2021, 118, e2101691118. [Google Scholar] [CrossRef]
  83. Bleibtreu, A.; Clermont, O.; Darlu, P.; Glodt, J.; Branger, C.; Picard, B.; Denamur, E. The rpoS gene is predominantly inactivated during laboratory storage and undergoes source-sink evolution in Escherichia coli species. J. Bacteriol. 2014, 196, 4276–4284. [Google Scholar] [CrossRef]
  84. Jiang, S.; Halbleib, A.; Antonovics, J.; Remus-Emsermann, M.; Rillig, M.C.; Mansour, I. Rapid evolution in necromass use under resource limitation reduces persistence in producer-decomposer microbial biospheres. Commun. Biol. 2026, 9, 755. [Google Scholar] [CrossRef]
  85. Arahal, D.R.; Gutierrez, M.C.; Volcani, B.E.; Ventosa, A. Taxonomic analysis of extremely halophilic archaea isolated from 56-years-old Dead sea brine samples. Syst. Appl. Microbiol. 2000, 23, 376–385. [Google Scholar] [CrossRef]
  86. Naidoo, G. Differential effects of nitrogen and phosphorus enrichment on growth of dwarf Avicennia marina mangroves. Aquat. Bot. 2009, 90, 184–190. [Google Scholar] [CrossRef]
  87. Lever, M.A.; Rogers, K.L.; Lloyd, K.G.; et al. Life under extreme energy limitation: a synthesis of laboratory- and field-based investigations. FEMS Microbiol. Rev. 2015, 39, 688–728. [Google Scholar] [CrossRef]
  88. Fendrihan, S.; Dornmayr-Pfaffenhuemer, M.; Gerbl, F.W.; et al. Spherical particles of halophilic archaea correlate with exposure to low water activity—implications for microbial survival in fluid inclusions of ancient halite. Geobiology 2012, 10, 424–433. [Google Scholar] [CrossRef]
  89. Setlow, P.; Kornberg, A. Biochemical studies of bacterial sporulation and germination: XXII. Energy metabolism in early stages of germination of Bacillus megaterium spores. J. Biol. Chem. 1970, 245, 3637–3644. [Google Scholar] [CrossRef]
  90. Adamski, J.C.; Roberts, J.A.; Goldstein, R.H. Entrapment of bacteria in fluid inclusions in laboratory-grown halite. Astrobiology 2006, 6, 552–562. [Google Scholar] [CrossRef]
  91. Elabed, H.; González-Tortuero, E.; Ibacache-Quiroga, C.; et al. Seawater salt-trapped Pseudomonas aeruginosa survives for years and gets primed for salinity tolerance. BMC Microbiol. 2019, 19, 142. [Google Scholar] [CrossRef]
  92. Schubert, B.A.; Timofeeff, M.N.; Lowenstein, T.K.; Polle, J.E. Dunaliella cells in fluid inclusions in halite: significance for long-term survival of prokaryotes. Geomicrobiol. J. 2010, 27, 61–75. [Google Scholar] [CrossRef]
  93. Sankaranarayanan, K.; Timofeeff, M.N.; Spathis, R.; Lowenstein, T.K.; Lum, J.K. Ancient microbes from halite fluid inclusions: optimized surface sterilization and DNA extraction. PLoS ONE 2011, 6, e20683. [Google Scholar] [CrossRef]
  94. Price, P.B.; Sowers, T. Temperature dependence of metabolic rates for microbial growth, maintenance, and survival. Proc. Natl. Acad. Sci. U S A 2004, 101, 4631–4636. [Google Scholar] [CrossRef]
  95. Kempes, C.P.; van Bodegom, P.M.; Wolpert, D.; Libby, E.; Amend, J.; Hoehler, T. Drivers of bacterial maintenance and minimal energy requirements. Front Microbiol. 2017, 8, 31. [Google Scholar] [CrossRef]
  96. Tijhuis, L.; Loosdrecht, M.C.; Heijnen, J.J. A thermodynamically based correlation for maintenance gibbs energy requirements in aerobic and anaerobic chemotrophic growth. Biotechnol. Bioeng. 1993, 42, 509–519. [Google Scholar] [CrossRef]
  97. Marschall, E.; Jogler, M.; Henßge, U.; Overmann, J. Large-scale distribution and activity patterns of an extremely low-light-adapted population of green sulfur bacteria in the Black Sea. Environ. Microbiol. 2010, 12, 1348–1362. [Google Scholar] [CrossRef]
  98. Legat, A.; Denner, E.B.; Dornmayr-Pfaffenhuemer, M.; et al. Properties of Halococcus salifodinae, an isolate from Permian rock salt deposits, compared with halococci from surface waters. Life 2013, 3, 244–259. [Google Scholar] [CrossRef]
  99. Ochman, H.; Wilson, A.C. Evolution in bacteria: evidence for a universal substitution rate in cellular genomes. J. Mol. Evol. 1987, 26, 74–86. [Google Scholar] [CrossRef]
  100. Moran, N.A.; Munson, M.A.; Baumann, P.; Ishikawa, H. A molecular clock in endosymbiotic bacteria is calibrated using the insect hosts. Proc. R Soc. B Biol. Sci. 1993, 253, 167–171. [Google Scholar] [CrossRef]
  101. Guixa-Boixereu, N.; Lysnes, K.; Pedrós-Alió, C. Viral lysis and bacterivory during a phytoplankton bloom in a coastal water microcosm. Appl. Environ. Microbiol. 1999, 65, 1949–1958. [Google Scholar] [CrossRef]
  102. Robinson, J.L.; Pyzyna, B.; Atrasz, R.G.; et al. Growth kinetics of extremely halophilic archaea (family halobacteriaceae) as revealed by arrhenius plots. J. Bacteriol. 2005, 187, 923–929. [Google Scholar] [CrossRef]
  103. Zhang, Q.; Chen, Y.; Shi, H. Bacterial strategies in prolonged stationary phase: tradeoff between cell growth, maintenance, and recycling. Phys. Rev. Res. 2023, 5, 013119. [Google Scholar] [CrossRef]
  104. Gefen, O.; Fridman, O.; Ronin, I.; Balaban, N.Q. Direct observation of single stationary-phase bacteria reveals a surprisingly long period of constant protein production activity. Proc. Natl. Acad. Sci. U S A 2014, 111, 556–561. [Google Scholar] [CrossRef]
  105. Greening, C.; Carere, C.R.; Rushton-Green, R.; et al. Persistence of the dominant soil phylum Acidobacteria by trace gas scavenging. Proc. Natl. Acad. Sci. U S A 2015, 112, 10497–10502. [Google Scholar] [CrossRef]
  106. Jaishankar, J.; Srivastava, P. Molecular basis of stationary phase survival and applications. Front Microbiol. 2017, 8, 2000. [Google Scholar] [CrossRef]
  107. Oren, A. Thymidine incorporation in saltern ponds of different salinities: estimation of in situ growth rates of halophilic archaeobacteria and eubacteria. Microb. Ecol. 1990, 19, 43–51. [Google Scholar] [CrossRef]
  108. Pedrós-Alió, C.; Calderón-Paz, J.I.; MacLean, M.H.; Medina, G.; Marrasé, C.; Gasol, J.M.; Guixa-Boixereu, N. The microbial food web along salinity gradients. FEMS Microbiol. Ecol. 2000, 32, 143–155. [Google Scholar] [CrossRef]
  109. Kucukyildirim, S.; Behringer, M.; Williams, E.M.; Doak, T.G.; Lynch, M. Estimation of the genome-wide mutation rate and spectrum in the archaeal species Haloferax volcanii. Genetics 2020, 215, 1107–1116. [Google Scholar] [CrossRef]
  110. Kucukyildirim, S.; Ozdemirel, H.O.; Lynch, M. Similar mutation rates but different mutation spectra in moderate and extremely halophilic archaea. G3 2023, 13, jkac303. [Google Scholar] [CrossRef]
  111. Conover, J.L.; Wendel, J.F. Deleterious mutations accumulate faster in allopolyploid than diploid cotton (Gossypium) and unequally between subgenomes. Mol. Biol. Evol. 2022, 39, msac024. [Google Scholar] [CrossRef]
  112. Wang, X.; Morton, J.A.; Pellicer, J.; Leitch, I.J.; Leitch, A.R. Genome downsizing after polyploidy: mechanisms, rates and selection pressures. Plant J. 2021, 107, 1003–1015. [Google Scholar] [CrossRef]
  113. Luiselli, J.; Rouzaud-Cornabas, J.; Lartillot, N.; Beslon, G. Genome streamlining: effect of mutation rate and population size on genome size reduction. Genome Biol. Evol. 2024, 16, evae250. [Google Scholar] [CrossRef]
  114. Mackwan, R.R.; Carver, G.T.; Drake, J.W.; Grogan, D.W. An unusual pattern of spontaneous mutations recovered in the halophilic archaeon Haloferax volcanii. Genetics 2007, 176, 697–702. [Google Scholar] [CrossRef]
  115. McGenity, T.J.; Gemmell, R.T.; Grant, W.D.; Stan-Lotter, H. Origins of halophilic microorganisms in ancient salt deposits. Environ. Microbiol. 2000, 2, 243–250. [Google Scholar] [CrossRef]
  116. Grant, W.D.; Gemmell, R.T.; McGenity, T.J. Halobacteria: the evidence for longevity. Extremophiles 1998, 2, 279–287. [Google Scholar] [CrossRef]
  117. Stan-Lotter, H.; Radax, C.; McGenity, T.J.; et al. From intraterrestrials to extraterrestrials—viable haloarchaea in ancient salt deposits. In Halophilic Microorganisms; Ventosa, A., Ed.; Springer: Berlin, 2004; pp. 89–102. [Google Scholar] [CrossRef]
  118. Fish, S.A.; Shepherd, T.J.; McGenity, T.J.; Grant, W.D. Recovery of 16S ribosomal RNA gene fragments from ancient halite. Nature 2002, 417, 432–436. [Google Scholar] [CrossRef]
  119. Maughan, H.; Birky, C.W.; Nicholson, W.L.; Rosenzweig, W.D.; Vreeland, R.H. The paradox of the “ancient” bacterium which contains “modern” protein-coding genes. Mol. Biol. Evol. 2002, 19, 1637–1639. [Google Scholar] [CrossRef]
  120. Speranza, G.; Cosentino, D.; Tecce, F.; Faccenna, C. Paleoclimate reconstruction during the Messinian evaporative drawdown of the Mediterranean Basin: Insights from microthermometry on halite fluid inclusions. Geochem Geophys Geosyst 2013, 14, 5054–5077. [Google Scholar] [CrossRef]
  121. Mansouri, H.; Prior, D.J.; Ajalloeian, R.; Elyaszadeh, R. Deformation and recrystallization mechanisms inferred from microstructures of naturally deformed rock salt from the diapiric stem and surface glaciers of a salt diapir in Southern Iran. J. Struct. Geol. 2019, 121, 10–24. [Google Scholar] [CrossRef]
  122. O'Neil, J.R.; Johnson, C.M.; White, L.D.; Roedder, E. The origin of fluids in the salt beds of the Delaware Basin, New Mexico and Texas. Appl. Geochem. 1986, 1, 265–271. [Google Scholar] [CrossRef]
Table Chronological list of reports of microbiological isolations from ancient halite.
Table Chronological list of reports of microbiological isolations from ancient halite.
Site Age Organisms Surface sterilization and culture methods Reference
Carey Salt Mine, Hutchinson, KS Permian
277-283 Myr		 Gram-positive halophilic diplococcus 70% alcohol for 15 sec [26]
Irkutsk, NY Precambrian 650 Myr 2 unidentified species Large crystal in Bunsen burner flame for 45 seconds. Drop into liquid media to dissolve till broth was saturated with NaCl. Tested with Pyocyaneus (Pseudomonas aeruginosa) [28]
Meyers, NY Silurian 3 unidentified species Large crystal in Bunsen burner flame for 45 seconds. Drop into liquid media to dissolve till broth was saturated with NaCl. Tested with Pyocyaneus (Pseudomonas aeruginosa) [28]
Saskatchewan, Canada Devonian 6 unidentified species Large crystal in Bunsen burner flame for 45 seconds. Drop into liquid media to dissolve till broth was saturated with NaCl. Tested with Pyocyaneus (Pseudomonas aeruginosa) [28]
Zechstein halite, Germany Permian
252.5-258 Myr		 “Bacillus circulans”
“Pseudomonas halocrenaea” 		Large crystal in Bunsen burner flame for 45 seconds. Drop into liquid media to dissolve till broth was saturated with NaCl. Tested with Pyocyaneus (Pseudomonas aeruginosa) [28,29]
Zechstein halite,
Germany		 Permian
252.5-258 Myr		 Mostly Bacillaceae members, other rods, Gram positive diplococci 1 mg/L Cl2 for 30 minutes, 80% ethanol for 30 minutes, or same as Dombrowski [31]
Winsford salt mine, Cheshire, England and Boulby potash mine, Cleveland, England Triassic Period, 195-225 Myr; Permian Period, 225-270 Myr Hbt. saccharovorum Hbt. salinarum
Haloarcula
Halococcus		 Newly exposed crystals submersion in absolute ethanol for 4-5 hours and dissolved in media, but other samples were untreated. [32]
Bad Ischl salt mine, Austria Permian Period, 225-280 Myr Hcc. Salifodinae
Halobacterium 		Dipping in ethanol and flaming; dissolved in media [34,33]
Salado Formation, New Mexico Permian Period, 250 Myr Virgibacillus 10 M NaOH, 10 M HCl, 5 min each; inclusion extracted with syringe [1]
Bad Ischl-Perneck, Austria; Altaussee, Austria; Berchtesgaden, Germany Permo-Triassic Period, 245-280 Myr Hcc. salifodinae Flaming with a Bunsen burner and monitoring temperature with an infrared thermometer; crystals dissolved [36]
Bad Ischl salt mine, Austria

Permian Period, 225-280 Myr Hcc. dombrowskii Flaming with a Bunsen burner and monitoring temperature with an infrared thermometer; crystals dissolved [37]
Death Valley, California Pleistocene Epoch, 97 Kyr Hbt. salinarum Inclusions were extracted using aseptic microdrill/micropipette after ethanol treatment [39]
Altaussee, Austria Permian Period, 225-280 Myr Hbt. noricense Flaming with a Bunsen burner and monitoring temperature with an infrared thermometer; crystals dissolved [38]
Sergipe Basin, Brazil Cretaceous Period, 121-122 Myr Halobacterium
Natronobacterium		 10 M NaOH, 10 M HCl, 5 min each; crystals dissolved in media [40]
Death Valley, California Pleistocene Epoch, 22-34 Kyr Halorubrum, Natronomonas, Haloterrigena 10 M NaOH for 5 min each; crystals dissolved in media [43]
Salar Grande, Chile Pliocene Epoch, 1.8-5.3 Myr Hbt. noricense 6% (w/v) sodium hypochlorite, 10 M NaOH, 10.53 M HCl, 70% (v/v) ethanol, 20 min each; crystals dissolved in media [42]
Penobsquis Mines, New Brunswick, Canada Mississippian, 350 Myr Multiple anaerobic curved rods/vibrio Rinsing in 70% alcohol, drying in UV hood for 10 minutes, extracting center portions, crushed and dissolved in anaerobic medium [47]
Yungying salt mine, China Eocene Epoch, 38-41 Myr Halobacterium, Halolamina 10 M NaOH, 10 M HCl, 5 minutes each; center of crystals chiseled out and dissolved [45]
Yunying Depression in Jianghan Basin of central China Eocene Epoch, 33.9-48.6 Myr Oceanobacillus 10 M NaOH, 10 M HCl, 10 min each; crushed with mortar and pestle [46]
Qianjiang Depression, China Permian Period, 123 Myr Hb. hubeinese 10 M NaOH, 10 M HCl, 5 minutes each; center of crystals chiseled out and dissolved [4]
Table 2. Quantitative Estimates Relevant to Microbial Survival Over Geological Timescales.
Table 2. Quantitative Estimates Relevant to Microbial Survival Over Geological Timescales.
Process or observation System Reported or calculated timescale Type of estimate Interpretive limitation
40K radiation model [58] Halophilic bacterial spores in a modeled halite fluid inclusion 17–34 Myr at 7 g/L K; 55–109 Myr at 2.2 g/L K Measured radiation-inactivation constants + dose model Assumes no active repair during dormancy, a 10-µL inclusion, 4000 CFU/mL initially, radiation is only one damage process.
Permafrost DNA and respiration [63] Environmental bacterial communities in frozen sediments 4-kb DNA and respiration detected at 400–600 kyr; neither detected at 740 kyr Environmental molecular and respiration observations Supports persistence to ~0.5 Myr in the studied permafrost; does not define a universal bacterial or spore survival limit.
Peptide-bond hydrolysis [64] Amino acids in deep-sea sediments ~1–2 Myr hydrolysis half-life in calcareous sediments Geochemical molecular-diagenesis estimate Not a microbial viability measurement; hydrolysis may be slower in non-carbonate sediments and depends on matrix conditions.
Aspartate racemization [12] G. stearothermophilus spores Half-life of 40–421 yr at 0–15°C; ~220 yr at 4°C High-temperature experiment + kinetic extrapolation Predicted 50% mortality times; dependent on extrapolation and environmental chemistry.
Thermal-inactivation extrapolation [11] Thermophilic Bacillus-group spores Extrapolated D-values ~1.9 × 10⁹–1.9 × 10¹² yr at 25–40°C High-temperature inactivation kinetics extrapolated downward A cautionary counterexample: the author called the upper values nonsensical and noted that brine-specific data were lacking.
Cultivable spores in sediments [66,67,68] Thermophilic and heterotrophic endospore populations Half-life of ~330 yr; 350 yr; calculated ~277–533 yr Environmental viable/culturable abundance decay Population-level loss of viability/culturability; burial, germination, deposition, and ecology may contribute.
DPA-derived endospore abundance [69] Global marine-sediment dataset Older segment: concentration only halved across ~20 Myr Segmented regression of DPA-derived abundance vs. sediment age Not a measured viability half-life; cross-site abundance integrates heterogeneous depositional and thermal histories.
Illustrative halite population calculation (this review) Hypothetical entrapped spore population ~3.7 × 10⁶ spores/metric ton initially; <1 expected after ~11,600 yr Author calculation using a 533-yr half-life Assumes 4000 CFU/mL brine, 10% survival/representation after entrapment, 2% fluid volume, and constant first-order decay.
LTSP extinction modeling [82] 21 heterotrophic bacterial taxa in long-term stationary phase Modeled population extinction ~1–100,000 yr Laboratory mortality trajectories + population model Not a sealed halite inclusion and not direct observation over the extrapolated interval; persistence varies strongly among taxa.
Dunaliella-supported energy budget [92] Miniaturized prokaryote supported by glycerol from one Dunaliella cell ~12 Myr published estimate Energy-budget calculation Assumes a 10-µm Dunaliella cell, 5.5 M glycerol, a 10-fg-C prokaryote, and a specified repair-metabolism demand; strongly assumption-dependent.
Note: These estimates are not directly interchangeable. Some quantify loss of culturability or population abundance, others quantify molecular damage, radiation exposure, or energy supply, and several depend on extrapolation beyond experimentally observed timescales. The table therefore summarizes quantitative constraints and comparisons rather than a set of equivalent maximum survival ages. Taken together, the estimates show both the multiplicity of processes relevant to long-term persistence and the strong dependence of inferred timescale on mechanism, environment, and model assumptions.
Disclaimer/Publisher’s Note: The statements, opinions and data contained in all publications are solely those of the individual author(s) and contributor(s) and not of MDPI and/or the editor(s). MDPI and/or the editor(s) disclaim responsibility for any injury to people or property resulting from any ideas, methods, instructions or products referred to in the content.
Copyright: This open access article is published under a Creative Commons CC BY 4.0 license, which permit the free download, distribution, and reuse, provided that the author and preprint are cited in any reuse.