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Signs of Life: A Report Based on the April 2000 Workshop on Life Detection Techniques Session 3: Detecting Extant Life
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Signs of Life: A Report Based on the April 2000 Workshop on Life Detection Techniques This page in the original is blank.
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Signs of Life: A Report Based on the April 2000 Workshop on Life Detection Techniques X-RAY MICROSCOPY AND THE DETECTION OF LIFE Chris Jacobsen Department of Physics and Astronomy State University of New York, Stony Brook Abstract The microscopic investigation of possibly life-containing specimens can be greatly aided by looking not just at the morphology of a feature of interest, but at its chemistry as well. Soft x-ray microscopes are well suited to this task. The basic ideas of soft x-ray microscopes, and their application to organic material detection from sample masses as small as 10−17 g, are briefly outlined. Introduction In the search for life elsewhere in the solar system, we have some expectations about what life might look like: It will likely involve carbon chemistry, and a spatial segregation of organic compounds in a way that simple chemical processes would not permit. Instrumentation for bulk chemical measurements on robotic probes may well provide more than enough of a signature to convince the scientific community of life elsewhere, but further proof (and proof that might be especially convincing to the general public) could be provided if we could actually see something in a sample returned to Earth that we could somehow recognize as representing life, either with something that was recently living or in a fossil record. However, seeing something that looks familiar may not be enough. For example, early reports of the imaging of DNA in scanning tunneling microscopes (STMs) used highly ordered pyrolitic graphite (HOPG) substrate, and subsequent investigations showed that at least some of the features reported were not those of DNA at all but of artifacts on the HOPG surface.1 Similar care must be applied to the interpretation of scanning electron microscope (SEM) images of intriguing features; ideally, one would like to know not just what is the morphology of the object under examination, but what is its composition. Better yet, when looking at the composition, one would like to go beyond a map of the concentration of elements in the specimen, to know of their chemical binding states. How does one map the chemistry of a specimen? Simply recording the x-ray fluorescence signal in an energy-dispersive detector in an SEM is not sufficient; the intrinsic energy resolution for detecting x-rays of an energy E (in electron volts) in a silicon detector is about , giving an energy width of about 30 eV for carbon K x-rays. Since chemical binding energies are in the range of 1-5 eV, one will have no ability to determine the chemical binding state of carbon in such a system. However, there are several microscopes that can be used to look at chemical binding states of carbon compounds: Secondary ion mass spectroscopy microscopes can provide 50-nm spatial resolution mapping of molecular masses, but because the specimen is eroded in the microscope, one can examine a microscopic feature only once. Infrared microspectroscopy provides very good chemical state mapping of organic molecules based on their well-defined vibrational and rotational states; it does so at about 10-µm resolution. Visible-light microscopes can be used to obtain some information about the chemistry of nonfluorescing compounds at a spatial resolution of 200 to 1000 nm. Electron energy loss spectroscopy allows one to look at chemical binding states of 100-nm thin section specimens, with about 0.5-eV energy resolution in most instruments and a spatial resolution of 5 nm or better. X-ray microscopy allows one to look at somewhat thicker sections, with an improved energy resolution of about 0.1 eV and a spatial resolution of 30-50 nm. While each of these tools has its own set of capabilities and limitations, x-ray microscopy is especially well suited to the examination of possible microbial life if samples can be returned to Earth.
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Signs of Life: A Report Based on the April 2000 Workshop on Life Detection Techniques Soft X-ray Microscopy We outline some of the characteristics of soft x-ray microscopes. More complete discussions can be found elsewhere.2−5 Photons in the energy range of about 100 to 1000 eV are often called “soft” x-rays (though usage of the term varies a bit). In this energy range, x-rays have a wavelength of a few nanometers, so that the potential is there for very high spatial resolution in microscopes. More importantly, soft x-rays are well suited to providing high spatial resolution images of organic specimens. By operating at photon energies above ~289 eV, one has enough energy to remove inner shell electrons from carbon atoms by ionization. However, if the photon energy is kept below ~540 eV, one will not have enough energy to remove inner shell electrons from oxygen. A consequence is that one can image hydrated, organic specimens with very high concentration by operating in the “water window” energy range between about 289 and 540 eV.6,7 Furthermore, by operating at energies slightly below the ionization threshold of carbon, the inner shell electron can be excited to an unoccupied molecular orbital. This gives rise to pre-edge absorption resonances that go by the name of x-ray absorption near-edge structure (XANES) or near-edge x-ray absorption fine structure (NEXAFS). This same effect forms the basis for energy loss near-edge structure (ELNES) in electron energy loss spectroscopy (EELS), except that in EELS the near-edge signal lies upon a large background of various plural inelastic scattering events. The intrinsically higher signal-to-noise ratio of XANES spectroscopy means that one can generally obtain better chemical state mapping information with less radiation dose to the specimen.8,9 While x-ray illumination of a specimen can be used to generate photoelectrons from the outermost 100-nm specimen layer that are then imaged with electron optics,10 transmission imaging is more commonly used for chemical state mapping of features located not just on a specimen surface. Most soft x-ray transmission microscopes use Fresnel zone plates as the focusing optic. These zone plates operate as circular diffraction gratings, and their Rayleigh spatial resolution can be as good as 1.22 times the outermost zone width. The best zone plates now available have an outermost zone width of 20-30 nm,11−13 giving a Rayleigh resolution of 25 to 35 nm. When used in scanning transmission x-ray microscopes with grating monochromators and undulator radiation,14,15 this spatial resolution can be combined with spectroscopy at 0.1 to 0.2-eV energy resolution,16,17 thereby matching the intrinsic width of XANES resonances. With this approach of soft x-ray spectromicroscopy (Figure 1), one can determine the bonding state of organic materials at 1 percent or greater concentration in a 50 × 50 × 200 nm3 volume, or a feature mass of only 10−17 g. FIGURE 1. X-ray spectromicroscopy in biology. The image on the left was one of six taken at specific photon energies in the carbon XANES spectral region (absorption spectra of a protein standard, bovine serum albumin [BSA], and of DNA are shown on the right). From these images, maps of protein and DNA distributions in bull sperm were obtained as part of a study of protamine binding. SOURCE: X. Zhang, R. Balhorn, J. Mazrimas, and J. Kirz, “Mapping and Measuring DNA to Protein Ratios in Mammalian Sperm Head by XANES Imaging,” Journal Struct. Biol. 116:335-344, 1996.
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Signs of Life: A Report Based on the April 2000 Workshop on Life Detection Techniques With these microscopes, one can in favorable circumstances take a single image of an initially living specimen in fully hydrated conditions; however, the radiation dose delivered to the specimen as part of the imaging process is on the order of 106 Gy, which produces immediate changes in a room-temperature specimen. 18 The solution is to examine the specimen at liquid nitrogen temperatures, where the diffusion of irradiation-produced free radicals is essentially stopped; in this case, specimens can withstand radiation doses of about 1010 Gy before mass loss is observed.19,20 When the specimen is stable enough to allow the acquisition of multiple images, one can rotate the specimen within the depth of focus of a zone plate and obtain tomographic reconstructions of the three-dimensional nature of micrometer-size specimens.21-23 X-ray Microscopy for Life Detection Soft x-ray microscopes are being used for studies in biology24,25 and polymer science,26,27 among other fields.28 In Figure 2, we show the examination of a thin section of the ALH84001 meteorite where one can determine that the ratio of organic carbon to carbonates is significantly higher in the “rim” region. Soft x-ray microscopy studies on astrobiology specimens are only beginning,29,30 but the combination of high spatial resolution and chemical state sensitivity should prove quite powerful. In organic geochemistry, Cody et al. have carried out investigations on the diagenesis of organic matter through geological time, including wood, fossilized plants and wood, and coal.31 X-ray microscopy has been able to reveal that signatures of life, including the presence of carbohydrates and recognizable cell walls, can be unambiguously identified even in partially mineralized fossils in a way that other techniques cannot address. We therefore conclude that x-ray microscopy may prove useful in the study of returned samples to aid the search for life outside of Earth. FIGURE 2. Illustration of the use of soft x-ray spectromicroscopy for the study of a thin-section sample of the ALH84001 meteorite. The image shown was taken at a photon energy of 284.0 eV with a pixel size of 48 nm. Different analysis regions were selected (two are indicated by arrows) to highlight a background region within the specimen; the carbonaceous-rich rim, which is rich in polycyclic aromatic hydrocarbons (PAHs); and the porous and globule regions of the meteor chip. The corresponding absorption spectra indicate that the rim has a higher ratio of organic material to carbonate. SOURCE: C. Jacobsen, S. Wirick, G. Flynn, and C. Zimba, “Soft X-ray Spectroscopy from Image Sequences with Sub-100 nm Spatial Resolution, ” Journal Microsc. 197:173-184, 2000.
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Signs of Life: A Report Based on the April 2000 Workshop on Life Detection Techniques CHARACTERIZING THE INTACT MICROBE-MINERAL INTERFACE William W. Barker and Jillian F. Banfield Department of Geology and Geophysics University of Wisconsin, Madison Abstract A predictive model for mineralogical and textural biosignatures has been developed from a descriptive model of geomicrobial influences on silicate mineral weathering by lithobiontic microbial communities. Zone 1, represented by the upper thallus of lichens, is devoid of substratum-derived minerals. In the lower thallus, physical disruption of the mineral substratum by microbial communities delineates Zone 2 (direct biochemilithic). Complex mixtures of organic polymers and secondary phases coat extensively corroded mineral surfaces. Mineral weathering reactions in Zone 3 (indirect biochemilithic) occur within spaces too small for microbes, resemble physiochemical processes, and are accelerated by soluble organic compounds. Physiochemical weathering and unaltered primary silicates characterize Zone 4. Textures defined by detrital and authigenic minerals trapped in extracellular polymers, mineralization of cell walls and sheaths, and the presence of biominerals may generate recognizable mineralogical biosignatures within Zones 1 and 2. Distinctive etch patterns of mineral surfaces may comprise mineralogical biosignatures within Zones 1, 2, and 3. Sample preparation techniques designed to preserve the morphology, antigenicity, and mineralogy of these highly hydrated and complex samples for high-resolution electron microscopy are an important tool for evaluating these materials. Introduction An ideal biosignature should be widespread, easily recognized, and preservable in the planetologic record. Despite the inherent necessity for biosignatures to be non-“Earth centric,” important clues may be extracted through examination of elemental, isotopic, mineralogical, and textural heterogeneities that arise due to interactions between living entities and their mineralogical environment. In the foreseeable future, time, resources, and engineering constraints will limit the search for extraterrestrial life to sites within our own solar system and to the near surface of these targets, where it is likely that organic material is scarce and the bulk of the material is mineralogical in nature. A reasonable biosignatures program must likewise address these realities. Herein we describe the microbe-mineral environment and potential mineralogical biosignatures in the context of a model based upon electron microscopic characterization of the intact organic-inorganic interface between silicate mineral assemblages and lithobiontic microbial communities, and laboratory studies. Interactions in the rhizosphere, soils, and sediments share similarities with those at the lichen-mineral interface, so data gathered on microbe-mineral interactions in the context of mineralogical biosignature formation may apply widely. Zone Model of Mineralogical Biosignature Formation The lichen mineral interface, comprised of an extremely complex microbial community in contact with a limited mineral assemblage grading from fresh to deeply weathered, is an ideal microcosm for studying processes that ultimately result in mineralogical biosignatures. Based on several years of high-resolution electron microscopic characterization of the intact microbe-mineral interface and supporting laboratory investigations, we developed a descriptive model (Figure 1) of biogeochemical weathering, which we have here adapted to predict the potential for biosignature formation.32 In Zones 1 and 2 of lichens (which for the purposes of geomicro-or astrobiology are best considered biofilms), mineral particles derived from either the air (Zone 1) or the underlying rock (Zone 2) are in intimate contact with a diverse microbial community. Minerals range from almost unaltered, cleavage-bound fragments to highly corroded grains displaying etch pits on external surfaces and extensive internal porosity. In all cases, mineral fragments are coated and bound together by films composed of complex mixtures of organic polymers and secondary minerals (Figure 2).
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Signs of Life: A Report Based on the April 2000 Workshop on Life Detection Techniques FIGURE 1. A zone model of mineralogical biosignatures. FIGURE 2. A filamentous cyanobacterium (F) inside a thick sheath (S), coccoid cells (B), and a complex mixture of nanocrystalline clays and oxyhydroxide minerals bound in extracellular polymers (C) define the complex geochemical environment of biosignature formation. High-pressure cryofixed, freeze substituted zero loss energy-filtered transmission electron microscopy micrograph of ultramicrotomed thin section.
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Signs of Life: A Report Based on the April 2000 Workshop on Life Detection Techniques While surface-parallel layers of adsorbed cations and water on mineral surfaces probably provide nucleation sites for clays, textures and distribution of secondary minerals also are controlled in large part by extracellular organic polymers. In addition to trapping and binding allochthonous mineral particles, these act as substrates for precipitation and affect crystallization of clays on mineral surfaces and in the polysaccharide capsules surrounding microbial cells. Zone 3 occurs within the intact substratum beneath the lichen community and grades upward into the “direct biochemilithic zone.” However, the small size of openings precludes microbial colonization, so it is defined based upon the absence of direct contact between high molecular weight microbial polymers and mineral surfaces. While silicate mineral reactions in this zone resemble those seen in strictly physiochemical weathering, they differ from the dissolution and recrystallization mechanisms seen in Zones 1 and 2. Comparison with nearby uncolonized surfaces, which show no evidence of reaction, indicates appreciable enhancement of weathering due to downward percolation of solutions containing dissolved low molecular weight organic products, especially acids. These findings have been quantified though experimental studies. 33 Consequently, we term this the “indirect biochemilithic” zone. A fourth “physiochemilithic” zone results from abiotic weathering. It may be found at distance from the microbe-mineral interface (as in Figure 1) or in environments where microorganisms are not present. This zone represents a standard against which mineralogical biosignatures must be measured, in part to ensure that nonbiological explanations cannot be provided for chemical and textural patterns detected. Possible biosignatures applicable to our model identified through characterization and experimental studies of microbe-mineral interactions include the following: Entrained or authigenic reaction products (e.g., clays) in extracellular polymers; Mineralization of cell walls and sheaths; and Crystallographically controlled etching of mineral surfaces. Recommended Analytical Protocols A research program in which qualitative findings from field studies of natural materials are quantitatively pursued by laboratory experiments is recommended. Characterization of complex mixtures of nanocrystalline minerals, highly hydrated extracellular polymers of variable composition and structure, and a diverse microbial assemblage is required to understand microbe-mineral interactions leading to formation and preservation of mineralogical biosignatures. The size of the components and the scale of the heterogeneities dictate the use of high-resolution analytical scanning and transmission electron microscopy in combination with biological cytochemical methods. Specimen preparation techniques must rigorously avoid changes in colloidal chemistry, hydration state, and mineralogy, all of which drastically affect organic-mineral adsorption and, hence, textural relationships. Additionally, specimen preparation must provide superior ultrastructural and antigenic preservation while minimizing changes in elemental concentrations. Many important characterization techniques will provide new information about processes occurring at the microbe-mineral interface. For example, elemental and valence state mapping by X-ray methods34 and surface characterization by atomic force microscopy35 appear promising. Ultrarapid cryoimmobilization is a superior specimen preparation technique for both scanning electron microscopy (SEM) and transmission electron microscopy because it minimizes dehydration artifacts. Furthermore, fixation is instantaneous (as opposed to the tens of minutes to hours required for more conventional chemical fixation methods) and provides superior ultrastructural and antigenic preservation. The goal of all cryofixation techniques is to freeze the water component rapidly and prevent sample damage from ice crystal nucleation and growth resulting in vitreous ice.36,37 High-pressure cryofixation offers the potential for achieving this in large volumes, in some cases up to 1 mm3. The ability to produce vitreous ice conditions in samples of these dimensions makes this technique particularly useful for preserving samples containing intact microbial biofilm communities on colonized minerals. Once cryoimmobilized, samples can be stored indefinitely under liquid nitrogen and subjected to a wide variety of preparation and analytical techniques (Figure 3).
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Signs of Life: A Report Based on the April 2000 Workshop on Life Detection Techniques FIGURE 3. Recommended analytical flow chart for samples containing mixtures of microbes and minerals. Given the aforementioned constraints imposed by geobiological samples, freeze substitution appears to be an extremely useful preparative method for mineralogical biosignatures research. Resultant epoxy-embedded samples are amenable to standard ultramicrotomy. Thin sections can be examined at high resolution with immunocytochemical methods and microanalytical and structural techniques appropriate for mineralogical investigation. Freeze fracture and examination of partially freeze dried samples in a cryostage-equipped field emission gun SEM is a useful correlative technique for examining the hydrated textures and structures of extracellular polymers, expansible clays, and microorganisms. These data are necessary to better understand formation of biosignatures as a consequence of biologically induced precipitation, dissolution, enzymatic redox reactions, uptake and redistribution of nutrients and toxins, and elemental mobility arising from complexation of metal ions by organics. Acknowledgments This research is supported by the Department of Energy Basic Energy Sciences, the National Science Foundation, the National Aeronautics and Space Administration, and NASA's Astrobiology Institute.
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Signs of Life: A Report Based on the April 2000 Workshop on Life Detection Techniques SINGLE-POLYMER MODEL DETECTION USING NANOPORES Amit Meller Rowland Institute at Harvard Daniel Branton Department of Molecular and Cellular Biology Harvard University Abstract The transmembrane channel created by α-hemolysin can be used to detect single linear charged molecules. Detection relies on drawing the linear molecule into the channel, thus reducing or blocking the ionic current that moves through the open pore. Translocation of the charged macromolecule through the channel is driven by a bias that is applied across the channel-containing membrane. This detection method does not rely on specific interactions between the protein pore and the translocating molecule. Rather, it is based on modulating the open pore cross section and depends on the “bulkiness” of the linear molecule. Information about the exact chemical composition of the molecules is not required to perform the measurements. Therefore, this method may lend itself to general use with a broad spectrum of uncharacterized materials. We have shown that different types of DNA molecules can be discriminated based on the blockade level and duration they produce when forced to translocate through our pore. Although the basic mechanism responsible for the translocation process and the resulting blockade level is not fully understood, we think that these measurements constitute the first step toward a fast and low-cost polymer characterization method. Introduction The genetic code of the form of life we are familiar with is stored in DNA molecules. By analogy to computers, we can think of DNA as the “media” used for code storage. When we speak about new life forms we may ask for alternative or analogue media to the familiar DNA. Although the existence of “alternatives” remains unknown, we can nonetheless conceive suitable tools to examine unfamiliar genetic material. When developing these tools one would face two fundamental requirements: First, since we may encounter situations where only small samples of the material under test will be available, our detection method should be able to resolve single molecules. Second, we would require a generic detection method that does not rely on particular enzymatic reactions but rather relies on more general properties of the molecule such as its bulkiness or local density. For example, the common modern techniques used for DNA sequencing make use of a particular set of nucleotides and rely on specific interaction between these molecules and enzymes.38 This chemistry may or may not be compatible with the new genetic material we will need to probe. We have recently developed a novel single-molecule DNA or RNA detection method that does not rely on specific interactions or chemistry. We have shown that DNA and RNA molecules can be probed by monitoring the ionic current blockade they produce as they are drawn through a narrow pore.39,40 We embed a single α-hemolysin pore from Staphylococcus aureus in a lipid bilayer that separates two small containers. The inner dimension of the self-assembled α-hemolysin channel is comparable with the typical cross section of polynucleotides. The pore allows free motion of small electrolytes that are present in the buffer solution. Charged molecules such as DNA or RNA are drawn through the channel under the influence of an external electric field (see Figure 1a). When a single polynucleotide molecule enters the pores it blocks most of the otherwise unper-turbed ionic current, thus signaling its presence in the pore. Our method makes use of sensitive ionic current measurements. We use a commercial patch-clamp electrometer to apply a constant electric field across the membrane and measure the resulting current. When a DNA molecule enters our pore the current drops from its initial (“open pore”) value of about 116 pA (at 120 mV applied voltage, KCl concentration of 1 M, and 22.0°C) to about 14 pA (see Figure 1b). The current is restored to its initial value when the molecule exits from the pore. We define the translocation time duration, tD , as the time that the signal stays at its lower state. We have found that tD is proportional to the linear length of the probed molecule.
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Signs of Life: A Report Based on the April 2000 Workshop on Life Detection Techniques FIGURE 1. (a) Schematic view of the nanopore detection method. The nanopore is embedded in a membrane separating two small containers filled with electrolyte solution. Charged polymers such as DNA are drawn through the pore under the influence of an electric field. The arrow indicates the translocation direction. (b) Typical current trace showing two DNA translocation events when a 120-mV gradient is applied across a membrane containing the α-hemolysin channel. For each event, we measured the translocation duration, tT, and the normalized blockade level defined as The fractional blockade level, IB, is defined as the ratio of the average current at the translocation period to the average open pore current. We have found that tD and IB may be used to discriminate different types of DNA molecule “on the fly.” Polynucleotides Discrimination To illustrate the capability of our system to discriminate single polynucleotides we characterized the blockade signals produced as homopolymers containing cytosines (poly(dC)100) or adenines (poly(dA)100) translocated through an α-hemolysin channel. Each DNA molecule was characterized by the duration of the blockade it produced, tD, and the average blockade current, IB. These parameters were plotted on an event diagram in which each point represents a single translocation event (Figure 2a and the corresponding histograms on Figures 2b and 2c). The most prominent features of these plots are the following: The events corresponding to the two polymers, each cluster in well-separated regions; less than 1 percent of the poly(dA)100 events fall in the poly(dC)100 region and vice versa. Thus, discrimination between the two polymer types is readily achieved. The poly(dA)100 events separate into two groups. So too do the poly(dC)100 events, albeit the separation into two groups is not as clear for poly(dC)100 as it is for poly(dA)100. The two groups are evident as two peaks in the current histograms for each polymer type (Figure 2b).
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Signs of Life: A Report Based on the April 2000 Workshop on Life Detection Techniques microbial contamination at different levels of resolution. This approach could be used for the detection of very general classes of assembly area microbial contaminants. Quantification of Populations or Single Cells There are two basic formats for using phylogenetic probes to study the environmental distribution of microorganisms. These are hybridization to total rRNA recovered from environmental samples and hybridization to fixed whole cells for subsequent microscopic visualization and enumeration. The latter technique, fluorescent in situ hybridization (FISH), uses fluorescent dye-labeled probes for microscopic detection of cells that hybridize to the probe. Both methods are well developed and in routine application.161-163 Both methods avoid the biases associated with PCR amplification by taking advantage of the natural amplification of the rRNAs. There are generally thousands of copies of each rRNA per cell, although the number for slow-growing cells is not well defined. However, both detection formats are much less sensitive than PCR-based methods. The lower limit of detection for radio-labeled probes is approximately 103-104 cells,164 and although a single cell can be observed microscopically, enumeration of low-abundance samples is limited by the image analysis requirements previously discussed. The efficiency of sample recovery and cell lysis or permeabilization are also poorly defined variables. Functional Probes An important distinction is made between probes designed to identify phylogenetic or taxonomic groups, and probes designed to monitor specific metabolic functions. There has been considerable development of probes targeting genes encoding specific enzymes to evaluate specific chemical transformations or potential activity of environmental populations. These “functional” gene probes should have application in determining whether spacecraft contaminants harbor microbial phenotypes more likely to proliferate in nonterrestrial habitats (e.g., autotrophs). Some examples of traits, and corresponding genes, sufficiently conserved to be easily identified in environmental samples include genes for nitrogen fixation,165 Ni-Fe hydrogenase,166 CO2 fixation, and the dissimilatory sulfite reductase.167,168 Other Technical Issues The following technical issues are key considerations in detection. However, since relatively little explicit or comparative data is available, they are only briefly listed here: Time and cost of analysis; Confidence levels; Signal to noise—the ability to “see” life in a complex background matrix; Sensitivity; Specificity; Analytical framework; Destructive sampling versus replication; In situ analyses versus the requirement for sample collection; Efficiency of sample collection and possible associated loss of viability; and Witness plates-active versus passive collection. A Possible Role for Improved Culture- and Activity-Based Methods The NASA Standard Assay is based on standard culture-based enumeration of cells and spores. However, the current protocol requires extended incubation periods and may miss more than 99 percent of microorganisms. Although this and other standard culture-based methods are now recognized to be inadequate, there may be merit in exploring alternative culture-based methods to either enumerate or measure physiological activity. For example, a recent study of low-abundance ice-entrained bacteria (200-300 cells/ml) showed that at least some of the assemblage was viable as measured by respiration of 14C-labeled acetate and glucose.169
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Signs of Life: A Report Based on the April 2000 Workshop on Life Detection Techniques A ROBOTIC-PCR DETECTOR FOR DNA-BASED LIFE ON OTHER PLANETS Gary Ruvkun Harvard Medical School, Department of Molecular Biology, Massachusetts General Hospital Michael Finney MJ Research, 590 Lincoln St., Waltham, Massachusetts Walter Gilbert Department of Molecular and Cellular Biology, Harvard University George M. Church Department of Genetics, Harvard Medical School Abstract Key genetic components such as the ribosomal RNA genes are conserved in all living organisms on Earth. The polymerase chain reaction (PCR) with DNA primers corresponding to conserved elements in the ribosomal RNA genes is now used to amplify minute quantities of DNA from extreme Earth environment samples, allowing the detection and classification of life in those samples without need to isolate, culture, or grow the organisms in any way. This is currently our most sensitive detector of life on Earth. These same primers can be used to search for life on Mars that is related to life on Earth. Life on Mars would be ancestrally related to life on Earth if there was exchange of viable organisms between Mars and Earth, or more generally, common ancestry from a source outside of the solar system, something we call Panarchaea. If microbial life exists on Mars currently and is more abundant than about one cell per gram at the landing site, PCR is sensitive enough to detect the signature of life. This signature is the amplification of a 500 base-pair-long DNA fragment that can be sequenced in situ on Mars using simple gene array technology to determine its phylogenetic placement. PCR technology is very mature. A low power consumption, very light, robotic PCR thermal cycler could be sent to the surface of Mars to sample the soil for microbial life. Even though life has so far only been found on Earth, most strategies for the detection of life on other planets and moons assume independent evolution on those planets. General life detection strategies involve the detection of structural and informational polymers, of microscopic or macroscopic structures, or of chemical signatures of metabolism. It is worth noting, however, that these are not the tools currently used by molecular biologists to search for the most divergent life on Earth. First, the rate of metabolism for many of the most divergent organisms, and the numbers of individual organisms in many niches, is so low that such detection tests currently fail. For the detection of the most diverse microbial life on Earth, there is a much more sensitive and simple DNA-based search strategy that is essentially definitive. The exploration of the diversity of life on Earth has revealed key genetic components that are conserved in all free-living organisms. The most conserved of these genes are the ribosomal RNA genes, one class of which is called the 16S or 18S RNA genes.170 Ribosomal RNAs are the main structural and catalytic components of the ribosome, the molecular machine that translates RNA into proteins. Because of this universality, all organisms are thought to have inherited their ribosomal RNA gene (as well as hundreds of other genes that have drifted more significantly during the past 3 billion years) from a common ancestor. The root of that tree has been hypothesized to be an archaeal-like hyperthermophile.171 Within the approximately 1500 nucleotides of the ribosomal RNA gene, there are multiple 20-nucleotide segments that are exactly conserved between organisms as disparate as the single-celled extremophile Archaeae that live in Earth's crust and humans, or corn.172 These conserved elements in the ribosomal genes are our most sensitive detectors of life on Earth and are being used to drastically expand the known universe of life. The reason they are so sensitive is that the PCR with DNA primers corresponding to these absolutely conserved elements can be used to amplify any DNA species, without the need to isolate, culture, or grow the organsims in any way.
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Signs of Life: A Report Based on the April 2000 Workshop on Life Detection Techniques The polymerase chain reaction amplifies the number of copies of a specific region of DNA in order to produce enough DNA, for example, for DNA sequence analysis. In PCR, one must know the sequences that flank a given region, but one need not know the DNA sequence in between. In addition to the range of organisms that can be detected, the PCR approach has the added advantages of extreme sensitivity and almost trivial ease. PCR can often detect a single DNA double helix in a clean sample in a two-hour analysis. The biochemical processing of the sample can be as crude as a cheek swab from humans to agitation of a soil sample. The technology of PCR involves adding stable 20-nucleotide-long DNA primers, a stable enzyme and nucleotide monomers that function at 370 Kelvin, and a simple heat pump that thermally cycles 40 times in two hours to amplify DNA from a few strands to billions of identical DNA fragments. In the case of the ribosomal genes, the DNA primer 5' GTGCCAGCAG CCGCGGTAAT, which corresponds to nucleotides 601 to 621 of a ribosomal gene, and GGTGGTCCTG ACCTCGGAC 5', which corresponds to the base pairing complement of nucleotides 1213 to 1233, are added to a soil extract. These DNA primers pair with their complement on each DNA strand, even if there are only a few DNA molecules in a sample. When DNA polymerase and mononucleotides are added, that DNA strand is duplicated. If one repeats the thermal cycle with all the same components in the same tube, there will be eight strands. Twenty cycles produce one million copies of the original sequences. This PCR technology is standard in small labs all over the world. Currently, hundreds of research groups are using ribosomal RNA PCR primers to amplify samples isolated from a wide range of environments —for example, from marine water samples or extreme environments.173 In this way, vast numbers of new archaeal and bacterial species have been found in extreme temperature or in dehydrated or radioactive environments as well as in the most typical ocean sample. Interestingly, most of the life that is detected in these samples cannot be cultured, suggesting either very slow growth rates or very particular growth conditions not met in the laboratory. This also suggests that metabolic detection of life can be very problematic since the growth rates can be slow or nil in simple culture conditions. Since universal PCR primers from the ribosomal RNA gene can detect essentially all living organisms on Earth, these same primers can be used to search for life on Mars that is related to life on Earth. If such microbial life exists on Mars currently and is more abundant than about one cell per gram at the landing site, PCR is sensitive enough to detect the signature of life—amplification of DNA with particular DNA sequence features using these DNA primers. There are two conditions under which life on Mars should resemble life on Earth to the point of sharing ribosomal RNA sequences: exchange between Mars and Earth or, more generally, common ancestry from another source, perhaps even outside of the solar system. In either case, organisms (or perhaps the hardier spores that many microbes form) would have to survive the passage. Ultraviolet radiation damage appears to be easily shielded for example, by simple embedding in a rock, though survival for millions of years has not been addressed. 174,175 The ability of a crustal sample to escape Mars and land on Earth without sterilizing heating is demonstrated by ALH84001 martian meteorite. While the ALH84001 meteorite evidence for fossils is debated, this rock may not have been heated above the survival temperature of modern spores,176 though this too is debated.177 The flux of such meteoritic exchange between Mars and Earth would be much more intense at the heavy bombardment stage of the solar system, as well as in the period of sporadic collisions towards the end of that era. Thus, life in the solar system may have evolved on Earth, Mars, or another planet and exchanged between those planets. There is also a reasonable argument for life on Earth coming from outside the solar system, something we call Panarchaea. The fossil record of Earth shows that within a few hundred million years of the cessation of the bombardment phase (at 4.0 billion to 4.2 billion years ago), living organisms morphologically similar to contemporary stromatolites, a mat composed of billions of bacteria, had already become abundant (at 3.6 billion years ago).178 This observation is usually marshaled as evidence that the evolution of life from simple molecules to
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Signs of Life: A Report Based on the April 2000 Workshop on Life Detection Techniques simple organisms was rapid. In that 300 million to 500 million years, life had to evolve from simple organic molecules to a bacteria-like organism with DNA-based genetic material, a very complex ribosome, and many other “core” genetic components now shared between the disparate branches on the tree of life.179 A different interpretation of the appearance of bacteria-like organisms within a short period of time after the end of bombardment is that it may represent the time it took for preexisting microbes to colonize the virgin chemical gradients and reduced carbon substrates in Earth 's crust. If one accepts that evolution to microbes can occur in 500 million years, then there may have been planets with microbial life that evolved during the 8 billion years of galactic evolution that preceded the birth of the solar system. Since planets are thought to be common and the habitable zone for life has broadened to much of the solar system, life may have evolved on many planets. Once life evolved in any planetary system, ejection of that life by meteorite and comet impacts or by chaotic planetary orbit pertubation and ejection could allow exchange within and between solar systems.180 Of course, if extraterrestrial life seeded Earth, Mars might have been seeded by the same source. The ability of microbial life to survive such long transits in vacuum and high radiation strains credulity, though protection could be afforded if the ejecta was as large as a planet or asteroid.181 However, the improbability of the arrival of life on Earth from another source must be compared to the similarly improbable scenario that the isotopic record in ancient rocks implies: that life evolved on Earth within 500 million years from primordial soup to full-fledged microbes. A strength of the Panarcheal argument is that it makes clear predictions of what extraterrestrial life should look like. For both the case of exchange between Mars and Earth and the general case of exchange between planetary systems, life on Mars should share ribosomal RNA genes with DNA sequence homology to earthly ribosomal genes, and PCR will be able to detect them. Life on Mars related to life on Earth may have flourished a billion years ago, when water was more plentiful. PCR has been used to detect DNA in samples thousands of years old but not a million or a billion years old—we do not expect to detect microbial fossils with this procedure. But microbial life may still flourish in the very low temperature and low hydration of martian soils. Microbes are very adaptable: we find them growing in the extreme cold and dryness of Antarctica, as well as the extreme heat and pressure of Earth 's crust.182 Martian microbes may have solved the problem of life at 150 to 210 K with near zero water. In addition, the signs of massive recent water flows on Mars suggest more hospitable microbial niches in the not too distant past.183 Also, evidence for recent martian volcanism, suggests that temperatures may rise below the martian surface, especially near the sites of the most recent volcanism.184 And even if life flourishes only in isolated oases away from any landing site, the dust storms evident on Mars may blow recently alive or living microbial cells and/or spores into the landing site and be detectable with a technique as sensitive as PCR. The technology to amplify and detect the ribosomal RNA signature of life on Mars is simple and can be engineered into a lightweight and reliable automated form. PCR thermal cyclers involve very simple heating and cooling that can use just a few watts. The technology is very mature, with thousands of solid-state thermal cycling machines, which cost from $1,000 to $10,000 each, installed in small laboratories all over the world. Many of these machines are robotically controlled in genome centers for high-throughput genome sequencing. PCR can thus be done with very little weight, low power consumption, and very forgiving technology. The soil sample handling capabilities of a future Mars lander could deliver samples to a suspension and dilution station. Filtration would be used to sieve out microbial-sized particles (microbial size on Earth is remarkably uniform, across vast phylogenetic distances). A simple extraction procedure can release the contents of any cells and remove contaminants. Samples would then be mixed with reagents including PCR primers corresponding to the universal elements of ribosomal genes and thermally cycled. Bona fide ribosomal RNA genes would be expected to generate a PCR product of a size in a narrow range. The DNA fragment from the PCR reaction would be probed to a gene array bearing 100,000 small oligonucleotides representing the DNA sequence space for the ribosomal RNA gene, to remotely determine the DNA sequence. In this way, any organism detected on Mars, could be placed in or outside of known microbial phylogeny. The phylogenetic placement of the detected ribosomal RNA gene is important for a quite different reason. PCR is so powerful that contamination is a critical issue. A ribosomal RNA gene sequence corresponding to a known earthly clade (such as humans or any of the common bacteria that live on the skin) would be rejected.
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Signs of Life: A Report Based on the April 2000 Workshop on Life Detection Techniques Another key to ruling out contamination will be dilution of the soil samples followed by PCR analysis; any true martian organism should be found in proportion to the input soil amount. There are also a number of well-established procedures, such as chemical and ultraviolet treatments to eliminate DNA contamination. The fluid handling components to handle the soil samples and transit to the PCR module could be microfluidic and weigh less than 100 grams. The thermal cycler for amplification could weigh just a few hundred grams. In principle, the DNA sequence analysis chip and charge coupled device electronics to read the chip scanner could be similar in size to a portable CD player. Such a micromicrobial analysis machine would also encourage useful advances in portable terrestrial DNA analyses as spinoffs. Acknowledgment We thank Jonathan Lunine for helpful comments on the manuscript.
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Signs of Life: A Report Based on the April 2000 Workshop on Life Detection Techniques REFERENCES FOR PAPERS IN SESSION 3 1. C.R. Clemmer and T.P. Beebe, “Graphite: A Mimic for DNA and Other Biolmolecules in Scanning Tunneling Microscope Studies,” Science 251:640-642, 1991. 2. See, for example, A.G. Michette, Optical Systems for Soft X Rays, Plenum Press, New York, 1986. 3. See, for example, J. Kirz, C. Jacobsen, and M. Howells, “Soft X-ray Microscopes and Their Biological Applications, Q. Rev. Biophys. 28:33-130, 1995. 4. See, for example, J. Thieme, G. Schmahl, E. Umbach, and D. Rudolph (eds.), X-ray Microscopy and Spectromicroscopy, Springer-Verlag, Berlin, 1998. 5. See, for example, W. Meyer-Ilse, D.T. Attwood, and A. Warwick (eds.), X-ray Microscopy: Proceedings of the Sixth International Conference, Berkeley, California, 2-6 August 1999, American Institute of Physics (AIP), Melville, New York, 2000. 6. H. Wolter, “Spiegelsysteme Streifenden Einfalls als Abbildende Optiken für Röntgenstrahlen,” Ann. Phys. 10:94-114, 286, 1952. 7. D. Sayre, J. Kirz, R. Feder, D.M. Kim, and E.D. Spiller, “Potential Operating Region for Ultrasoft X-ray Microscopy of Biological Materials,” Science 196:1339-1340, 1977. 8. M. Isaacson and M. Utlaut, “A Comparison of Electron and Photon Beams for Determining Micro-chemical Environment,” Optik 50:213-234, 1978. 9. E.G. Rightor, A.P. Hitchcock, and H. Ade, “Spectromicroscopy of Polyethylene terephthalate: Comparison of Spectra and Radiation Damage Rates in X-ray Absorption and Electron Energy Loss,” Journal of Physical Chemistry B 101:1950-1960, 1997. 10. See, for example, G. De Stasio, D. Dunham, B.P. Tonner, D. Mercanti, M.T. Ciotti, P. Perfetti, and G. Margaritondo, “Application of Photoelectron Spectromicroscopy to a Systematic Study of Toxic and Natural Elements in Neurons,” Journal of Synchrotron Radiation 2:106-112, 1995. 11. S. Spector, C. Jacobsen, and D. Tennant, “Process Optimization for Production of Sub-20 nm Soft X-ray Zone Plates,” Journal of Vacuum Science and Technology B 15:2872-2876, 1997. 12. E. Anderson and D. Kern, “Nanofabrication of Zone Plate Lenses for High-resolution X-ray Microscopy, ” in X-ray Microscopy III: Proceedings of the Third International Conference, London, September 3-7, 1990, A.G. Michette, G.R. Morrison, and C.J. Buckley (eds.), Springer-Verlag, New York, 1992. 13. D. Weiss, M. Peuker, and G. Schneider, “Radiation-enhanced Network Formation in Copolymer Galvanoforms for Diffractive Nickel X-ray Optics with High Aspect Ratios,” Applied Physics Letters 72:1805-1807, 1998. 14. C. Jacobsen, S. Williams, E. Anderson, M.T. Browne, C.J. Buckley, D. Kern, J. Kirz, M. Rivers, and X. Zhang, “Diffraction-limited Imaging in a Scanning Transmission X-ray Microscope, ” Optics Communications 86:351-364, 1991. 15. T. Warwick, H. Ade, S. Cerasari, J. Denlinger, K. Franck, A. Garcia, S. Hayakawa, A. Hitchcock, J. Kikuma, S. Klingler, J. Kortright, G. Morisson, M. Moronne, E. Rightor, E. Rotenberg, S. Seal, H.-J. Shin, W.F. Steele, and B.P. Tonner, “Development of Scanning X-ray Microscopes for Materials Science Spectromicroscopy at the Advanced Light Source,” Journal of Synchrotron Radiation 5:1090-1092, 1998. 16. X. Zhang, H. Ade, C. Jacobsen, J. Kirz, S. Lindaas, S. Williams, and S. Wirick, “Micro-XANES: Chemical Contrast in the Scanning Transmission X-ray Microscope,” Nuclear Instruments and Methods in Physics Research A 347:431-435, 1994. 17. C. Jacobsen, S. Wirick, G. Flynn, and C. Zimba, “Soft X-ray Spectroscopy from Image Sequences with Sub-100 nm Spatial Resolution,” Journal Microsc. 197:173-184, 2000. 18. See, for example, J. Kirz, C. Jacobsen, and M. Howells, “Soft X-ray Microscopes and Their Biological Applications, Q. Rev. Biophys. 28:33-130, 1995. 19. G. Schneider, “Cryo X-ray Microscopy with High Spatial Resolution in Amplitude and Phase Contrast,” Ultramicroscopy 75:85-104, 1998. 20. J. Maser, A. Osanna, Y. Wang, C. Jacobsen, J. Kirz, S. Spector, B. Winn, and D. Tennant, “Soft X-ray Microscopy with a Cryo Scanning Transmission X-ray Microscope: I. Instrumentation, Imaging and Spectroscopy,” Journal Microsc. 197:68-79, 2000. 21. W.S. Haddad, I. McNulty, and J.E. Trebes, “Ultrahigh-resolution X-ray Tomography,” Science 266:1213-1215, 1994. 22. J. Lehr, “3D X-ray Microscopy: Tomographic Imaging of Mineral Sheaths of Bacteria Leptothrix ochracea with the Göttingen X-ray Microscope at BESSY,” Optik 104:166-170, 1997. 23. Y. Wang, C. Jacobsen, J. Maser, and A. Osanna, “Soft X-ray Microscopy with a Cryo Scanning Transmission X-ray Microscope: II. Tomography,” Journal Microsc. 197:80-93, 2000. 24. See, for example, Figure 1 from X. Zhang, R. Balhorn, J. Mazrimas, and J. Kirz, “Mapping and Measuring DNA to Protein Ratios in Mammalian Sperm Head by XANES Imaging,” Journal Struct. Biol. 116:335-344, 1996. 25. J. Kirz, C. Jacobsen, and M. Howells, “Soft X-ray Microscopes and Their Biological Applications, Q. Rev. Biophys. 28:33-130, 1995. 26. H. Ade, X. Zhang, and S.H. Cameron, “Chemical Contrast in X-ray Microscopy and Spatially Resolved XANES Spectroscopy of Organic Specimens,” Science 258:972-975, 1992. 27. H. Ade, “Compositional and Orientational Characterization of Polymeric Systems with X-ray Microscopy,” Trends in Polymer Science 5:58-66, 1997. 28. For further examples see, for example, W. Meyer-Ilse, D.T. Attwood, and A. Warwick (eds.), X-ray Microscopy: Proceedings of the Sixth International Conference, Berkeley, California, 2-6 August 1999, American Institute of Physics (AIP), Melville, New York, 2000. 29. C. Jacobsen, S. Wirick, G. Flynn, and C. Zimba, “Soft X-ray Spectroscopy from Image Sequences with Sub-100 nm Spatial Resolution,” Journal Microsc. 197:173-184, 2000.
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Signs of Life: A Report Based on the April 2000 Workshop on Life Detection Techniques 30. G.J. Flynn, L.P. Keller, C. Jacobsen, and S. Wirick, “Carbon and Potassium Mapping and Carbon Bonding State Measurements on Interplanetary Dust,” Meteoritics and Planetary Science 33(supplement):A50-51, 1998. 31. G. Cody, R.E. Botto, H. Ade, and S. Wirick, “Soft X-ray Microscopy and Microanalysis: Applications in Organic Geochemistry,” in X-ray Microbeam Technology and Applications, W. Yun (ed.), International Society for Optical Engineering (SPIE), Bellingham, Washington, 1995. 32. W.W. Barker and J.F. Banfield, “Zones of Chemical and Physical Interaction at Interfaces Between Microbial Communities and Minerals: A Model,” Geomicro. Journal 15:223-244, 1998. 33. S.A. Welch, W.W. Barker, and J.F. Banfield, “Microbial Extracellular Polysaccharides and Plagioclase Dissolution, ” Geochim. Cosmochim. Acta 63:1405-1409, 1999. 34. S.C.B. Myneni, J.T. Brown, G.A. Martinez, and W. Meyer-Ilse, “Imaging of Humic Substance Macromolecular Structures in Water and Soils,” Science 286:1335-1337, 1999. 35. H.H. Teng and P.M. Dove, “Surface Site-specific Interactions of Aspartate with Calcite During Dissolution: Implications for Biomineralization,” Am. Mineral. 82:878-887, 1997. 36. See P. Echlin, Low-Temperature Microscopy and Analysis, Plenum Press, New York, 1992, 539 pp. 37. See R.A. Steinbrecht and K. Zierold, Cryotechniques in Biological Electron Microscopy, Springer-Verlag, New York, 1987, 297 pp. 38. R.C. Cantor and L.C. Smith, Genomics: The Science and Technology Behind the Human Genome Project , John Wiley & Sons, New York, 1999. 39. J.J. Kasianowicz, E. Brandin, D. Branton, and D.W. Deamer, “Characterization of Individual Polynucleotide Molecules Using a Membrane Channel,” Proc. Natl. Acad. Sci. USA 93:13770-13773, 1996. 40. M. Akeson, D. Branton, J.J. Kasianowicz, E. Brandin, and D.W. Deamer, “Microsecond Time-scale Discrimination Among Polycytidylic Acid, Polyadenylic Acid, and Polyuridylic Acid as Homopolymers or as Segments Within Single RNA Molecules,” Biophys. J. 77:3227-3233, 1999. 41. A. Meller, L. Nivon, E. Brandin, J. Golovchenko, and D. Branton, “Rapid Nanopore Discrimination Between Single Polynucleotide Molecules, ” Proc. Natl. Acad. Sci USA. 97:1079-1084, 2000. 42. A. Meller, L. Nivon, E. Brandin, J. Golovchenko, and D. Branton, “Rapid Nanopore Discrimination Between Single Polynucleotide Molecules, ” Proc. Natl. Acad. Sci USA 97:1079-1084, 2000. 43. A. Meller, L. Nivon, and D. Branton, “Voltage-driven DNA Translocations Through a Nanopore,” Phys. Rev. Lett. 86:3435-3438, 2001. 44. G.A. Logan, J.M. Hayes, G.B. Hieshima, and R.E. Summons, “Terminal Proterozoic Reorganization of Biogeochemical Cycles,” Nature 376:53-56, 1995. 45. S. Ziegler and M.L. Fogel, “Seasonal and Diel Determinants of the Isotopic Composition of Organic Matter in a Freshwater Wetland,” American Chemical Society Abstract 221, GEOC 61, 2001. 46. See paper by D. Stahl, Session 3, in this appendix. 47. See paper by J.M. Muldowan, Session 4, in this appendix. 48. M.D. McCarthy, J.I. Hedges, and R. Benner, “Bacterial Origin of a Major Fraction of Dissolved Organic Nitrogen in the Sea,” Science 281:231-233, 1998. 49. M.L. Fogel and N. Tuross, “Transformation of Plant Biochemicals to Geological Macromolecules During Early Diagenesis,” Oecologia 120:336-346, 1999. 50. See paper by R.J. Cotter, Session 3, in this appendix. 51. E.J. Gaidos, K.H. Nealson, and J.L. Kirschvink, “Life in Ice-covered Oceans,” Science 284:1631-1633, 1999. 52. H.A. Lowenstam and S. Weiner, On Biomineralization, Oxford University Press, Oxford, U.K., 1989, p. 324. 53. J.F. Kasting, “Earth's Early Atmosphere,” Science 259:920-926, 1993. 54. D.A. Evans, N.J. Beukes, and J.L. Kirschvink, “Low-latitude Glaciation in the Paleoproterozoic,” Nature 386(6622):262-266, 1997. 55. J.L. Kirschvink, E.J. Gaidos, L.E. Bertani, N.J. Beukes, J. Gutzmer, L.N. Maepa, and R.E. Steinberger, “Paleoproterozoic Snowball Earth: Extreme Climatic and Geochemical Global Change and Its Biological Consequences,” Proc. Natl. Acad. Sci. USA 97:1400-1405, 2000. 56. R. Rye and H.D. Holland, “Paleosols and the Evolution of Atmospheric Oxygen: A Critical Review, ” Amer. Journal Sci. 298(8):621-672, 1998. 57. P.M. Harrison, “The Structure and Function of Ferritin,” Biochem. Ed. 14:154-162, 1986. 58. S. Mann, J. Hannington, and R. Williams, “Phospholipid Vesicles as a Model System for Biomineralization,” Nature 324:565-568, 1986. 59. J.L. Kirschvink and H.A. Lowenstam, “Mineralization and Magnetization of Chiton Teeth: Paleomagnetic, Sedimentologic, and Biologic Implications of Organic Magnetite,” Earth Planet. Sci. Lett. 44:193-204, 1979. 60. J.L. Kirschvink, “Paleomagnetic Evidence for Fossil Biogenic Magnetite in Western Crete, ” Earth Planet. Sci. Lett. 59:388-392, 1982. 61. H. Vali and J.L. Kirschvink, “Observations of Magnetosome Organization, Surface Structure, and Iron Biomineralization of Undescribed Magnetic Bacteria: Evolutionary Speculations,” in Iron Biomineralization, R.P. Frankel and R.P. Blakemore (eds.), Plenum Press, New York, 1991, pp. 97-115. 62. R.P. Blakemore, “Magnetotactic Bacteria,” Science 190:377-379, 1975. 63. R.B. Frankel, R.P. Blakemore, and R.S. Wolfe, “Magnetite in Freshwater Magnetotactic Bacteria,” Science 203:1355-1356, 1979.
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Signs of Life: A Report Based on the April 2000 Workshop on Life Detection Techniques 64. H. Vali and J.L. Kirschvink, “Observations of Magnetosome Organization, Surface Structure, and Iron Biomineralization of Undescribed Magnetic Bacteria: Evolutionary Speculations,” in Iron Biomineralization, R.P. Frankel and R.P. Blakemore (eds.), Plenum Press, New York, 1991, pp. 97-115. 65. Y.A. Gorby, T.J. Beveridge, and R.P. Blakemore, “Characterization of the Bacterial Magnetosome Membrane,” Journal Bacteriol. 170:834-841, 1988. 66. R. Blakemore, “Magnetotactic Bacteria,” Annu. Rev. Microbiol. 36:217-238, 1982. 67. R.P. Blakemore, “Magnetotactic Bacteria,” Science 190:377-379, 1975. 68. J.L. Kirschvink, “South-seeking Magnetic Bacteria,” Journal of Experimental Biology 86:345-347, 1980. 69. R.P. Blakemore, R.B. Frankel, and A.J. Kalmijn, “South-seeking Magnetotactic Bacteria in the Southern Hemisphere,” Nature 286:384-385, 1980. 70. R.B. Frankel, R.P. Blakemore, F.F. Torres de Araujo, E.M.S. Esquivel, and J. Danon, “Magnetotactic Bacteria at the Geomagnetic Equator,” Science 212:1269-1270, 1981. 71. S.-B.R. Chang and J.L. Kirschvink, “Magnetofossils, the Magnetization of Sediments, and the Evolution of Magnetite Biomineralization,” Annu. Rev. Earth Planet. Sci. 17:169-195, 1989. 72. F.F. Torres de Araujo, M.A. Pires, R.B. Frankel, and C.E.M. Bicudo, “Magnetite and Magnetotaxis in Algae,” Biophys. Journal 50:375-378, 1985. 73. H. Vali and J.L. Kirschvink, “Observations of Magnetosome Organization, Surface Structure, and Iron Biomineralization of Undescribed Magnetic Bacteria: Evolutionary Speculations,” in Iron Biomineralization, R.P. Frankel and R.P. Blakemore (eds.), Plenum Press, New York, 1991, pp. 97-115. 74. S. Mann, R.B. Frankel, and R.P. Blakemore, “Structure, Morphology and Crystal Growth of Bacterial Magnetite,” Nature 310:405-407, 1984. 75. S. Mann, T.T. Moench, and R.J.P. Williams, “A High-resolution Electron Microscopic Investigation of Bacterial Magnetite: Implications for Crystal Growth,” Proceedings of the Royal Society of London B 221:385-393, 1984. 76. S. Mann, “Structure, Morphology, and Crystal Growth of Bacterial Magnetite, ” in Magnetite Biomineralization and Magnetoreception in Animals: A New Biomagnetism, J.L. Kirschvink, D.S. Jones, and B.J. McFadden (eds.), Plenum Press, New York, 1985, pp. 311-332. 77. H. Vali and J.L. Kirschvink, “Observations of Magnetosome Organization, Surface Structure, and Iron Biomineralization of Undescribed Magnetic Bacteria: Evolutionary Speculations,” in Iron Biomineralization, R.P. Frankel and R.P. Blakemore (eds.), Plenum Press, New York, 1991, pp. 97-115. 78. J.L. Kirschvink, “Magnetite Biomineralization and Geomagnetic Sensitivity in Higher Animals: An Update and Recommendations for Future Study,” Bioelectromagnetics 10:239-259, 1989. 79. J.L. Kirschvink, “On the Magnetostatic Control of Crystal Orientation and Iron Accumulation in Magnetosomes,” Automedica 14:257-269, 1992. 80. R.F. Butler and S.K. Banerjee, “Theoretical Single-domain Size Range in Magnetite and Titanomagnetite, ” Journal of GeophysicalResearch 80:4049-4058, 1975. 81. J.C. Diaz-Ricci and J.L. Kirschvink, “Magnetic Domain State and Coercivity Predictions for Biogenic Greigite (Fe3O4): A Comparison of Theory with Magnetosome Observations,” Journal of Geophysical Research 97:17309-17315, 1992. 82. K.L. Thomas-Keprta et al., “Elongated Prismatic Magnetite Crystals in ALH84001 Carbonate Globules: Potential Martian Magnetofossils,” Geochim. Cosmochim. Acta 64:6049-6081, 2000. 83. J.L. Kirschvink and S.-B.R. Chang, “Ultra Fine-grained Magnetite in Deep-sea Sediments: Possible Bacterial Magnetofossils,” Geology 12:559-562, 1984. 84. N. Petersen, T. von Dobeneck, and H. Vali, “Fossil Bacterial Magnetite in Deep-sea Sediments from the South Atlantic Ocean,” Nature 320:611-615, 1986. 85. J.L. Gould, J.L. Kirschvink, and K.S. Deffeyes, “Bees Have Magnetic Remanence,” Science 201:1026-1028, 1978. 86. C. Walcott, J.L. Gould, and J.L. Kirschvink, “Pigeons Have Magnets,” Science 205:1027-1029, 1979. 87. J.L. Kirschvink and J.L. Gould, “Biogenic Magnetite as a Basis for Magnetic Field Detection in Animals, ” Biosystems 13:181-201, 1981. 88. R. Wiltschko and W. Wiltschko, “Magnetic Orientation in Animals,” Zoophysiology, Vol. 33, Springer, Berlin, 1995, p. 297. 89. M.M. Walker, C.E. Diebel, and C.V. Haugh, “Structure and Function of the Vertebrate Magnetic Sense,” Nature 390:371-376, 1997. 90. J.L. Kirschvink, “Magnetoreception: Homing in on Vertebrates,” Nature 390:339-340, 1997. 91. M.M. Walker, C.E. Diebel, and C.V. Haugh, “Structure and Function of the Vertebrate Magnetic Sense,” Nature 390:371-376, 1997. 92. C.E. Diebel, R. Proksch, C.R. Green, P. Neilson, and M.M. Walker, “Magnetite Defines a Vertebrate Magnetoreceptor,” Nature 406:299-302, 2000. 93. J.L. Kirschvink and A. Kobayashi-Kirschvink, “Is Geomagnetic Sensitivity Real? Replication of the Walker-Bitterman Conditioning Experiment in Honey Bees,” American Zoologist 31:169-185, 1991. 94. J.L. Kirschvink, S. Padmanabha, C.K. Boyce, and J. Oglesby, “Measurement of the Threshold Sensitivity of Honeybees to Weak, Extremely Low Frequency Magnetic Fields,” Journal of Experimental Biology 200:1363-1368, 1997. 95. R.C. Beason, R. Wiltschko, and W. Wiltschko, “Pigeon Homing: Effects of Magnetic Pulses on Initial Orientation, ” Auk 114(3):405-415, 1997. 96. U. Munro, J.A. Munro, J.B. Phillips, and W. Wiltschko, “Effect of Wavelength of Light and Pulse Magnetization on Different Magnetoreception Systems in a Migratory Bird,” Australian Journal of Zoology 45(2):189-198, 1997.
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Signs of Life: A Report Based on the April 2000 Workshop on Life Detection Techniques 97. U. Munro, J.A. Munro, J.B. Phillips, R. Wiltschko, and W. Wiltschko, “Evidence for a Magnetite-based Navigational Map in Birds,” Naturwissenschaften 84(1):26-28, 1997. 98. W. Wiltschko, U. Munro, R.C. Beason, H. Ford, and R. Wiltschko, “A Magnetic Pulse Leads to a Temporary Deflection in the Orientation of Migratory Birds,” Experientia 50:697-700, 1994. 99. W. Wiltschko and R. Wiltschko, “Migratory Orientation of European Robins Is Affected by the Wavelength of Light as Well as by a Magnetic Pulse,” Journal of Comparative Physiology 177(3):363-369, 1995. 100. J.L. Kirschvink and S.-B.R. Chang, “Ultra Fine-grained Magnetite in Deep-sea Sediments: Possible Bacterial Magnetofossils,” Geology 12:559-562, 1984. 101. S.-B.R. Chang and J.L. Kirschvink, “Magnetofossils, the Magnetization of Sediments, and the Evolution of Magnetite Biomineralization,” Annu. Rev. Earth Planet. Sci. 17:169-195, 1989. 102. J.L. Kirschvink and H.A. Lowenstam, “Mineralization and Magnetization of Chiton Teeth: Paleomagnetic, Sedimentologic, andBiologic Implications of Organic Magnetite,” Earth Planet. Sci. Lett. 44:193-204, 1979. 103. S.-B.R. Chang and J.L. Kirschvink, “Magnetofossils, the Magnetization of Sediments, and the Evolution of Magnetite Biomineralization,” Annu. Rev. Earth Planet. Sci. 17:169-195, 1989. 104. J.L. Kirschvink and S.-B.R. Chang, “Ultra Fine-grained Magnetite in Deep-sea Sediments: Possible Bacterial Magnetofossils,” Geology 12:559-562, 1984. 105. H. Vali, O. Forster, G. Amarantidis, and N. Petersen, “Magnetotactic Bacteria and Their Magnetofossils in Sediments,” Earth Planet. Sci. Lett. 86:389-400, 1987. 106. T. von Dobeneck, N. Petersen, and H. Vali, “Bakterielle Magnetofossilien,” Geowissenschaften in unser Zeit 5:27-35, 1987. 107. H. Vali and J.L. Kirschvink, “Magnetofossil Dissolution in a Paleomagnetically Unstable Deep-sea Sediment,” Nature 339:203-206, 1989. 108. B.M. Moskowitz, R.B. Frankel, and D.A. Bazylinski, “Rock Magnetic Criteria for the Detection of Biogenic Magnetite,” Earth Planet. Sci. Lett. 120(3-4):283-300, 1993. 109. P.P. Hesse, “Evidence for Bacterial Paleoecological Origin of Mineral Magnetic Cycles in Oxic and Sub-oxic Tasman Sea Sediments,” Marine Geology 117:1-17, 1994. 110. T. Yamazaki and H. Kawahata, “Organic Carbon Flux Controls the Morphology of Magnetofossils in Marine Sediments,” Geology 16(12):1064-1066, 1998. 111. Z. Gibbs-Eggar, “Possible Evidence for Dissimilatory Bacterial Magnetite Dominating the Magnetic Properties of Recent Lake Sediments,” Earth Planet. Sci. Lett. 168:1-6, 1999. 112. R.B. Frankel, J.P. Zhang, and D.A. Bazylinski, “Single Magnetic Domains in Magnetotactic Bacteria, Journal of Geophysical Research 103:30601-30604, 1998. 113. J.A. Peck and J.W. King, “Magnetofossils in the Sediment of Lake Baikal, Siberia,” Earth Planet. Sci. Lett. 140(1-4):159-172, 1996. 114. P.P. Hesse, “Evidence for Bacterial Paleoecological Origin of Mineral Magnetic Cycles in Oxic and Sub-oxic Tasman Sea Sediments, Marine Geology 117:1-17, 1994. 115. J.F. Stolz, “Magnetosomes,” Journal of General Microbiology 139:1663-1670, 1993. 116. T. Yamazaki, I. Katsura, and K. Marumo, “Origin of Stable Remanent Magnetization of Siliceous Sediments in the Central Equatorial Pacific,” Earth Planet. Sci. Lett. 105(1-3):81-93, 1991. 117. J. Akai, T. Sato, and S. Okusa, “TEM Study on Biogenic Magnetite in Deep-sea Sediments from the Japan Sea and the Western Pacific Ocean,” Journal of Electron Microscopy 40(2):110-117, 1991. 118. M. Farina, D.M.S. Esquivel, and H. Debarros, “Magnetic Iron-sulfur Crystals from a Magnetotactic Microorganism, ” Nature 343(6255):256-258, 1990. 119. S. Mann, N.H.C. Sparks, R.B. Frankel, D.A. Bazylinski, and H.W. Jannasch, “Biomineralization of Ferrimagnetic Greigite (Fe3S4) and Iron Pyrite (FeS2) in a Magnetotactic Bacterium,” Nature 343:258-261, 1990. 120. D.A. Bazylinski, R.B. Frankel, and H.W. Jannasch, “Anaerobic Magnetite Production by a Marine, Magnetotactic Bacterium, ” Nature 334:518-519, 1988. 121. D.S. McKay, E.K. Gibson, K.L. Thomas-Krepta, H. Vali, C.S. Romanek, S.J. Clemett, X.D.F. Chillier, C.R. Maechling, and R.N. Zare, “Search for Past Life on Mars: Possible Relic Biogenic Activity in Martian Meteorite ALH84001,” Science 273:924-930, 1996. 122. K.L. Thomas-Keprta et al., “Elongated Prismatic Magnetite Crystals in ALH84001 Carbonate Globules: Potential Martian Magnetofossils,” Geochim. Cosmochim. Acta 64:4049-4081, 2000. 123. L.E. Borg, J.N. Connelly, L.E. Nyquist, C.-Y. Shih, H. Wiesmann, and Y. Reese, “The Age of the Carbonates in Martian Meteorite ALH84001,” Science 286:90-94, 1999. 124. B.P. Weiss, J.L. Kirschvink, F.J. Baudenbacher, H. Vali, N.T. Peters, F.A. MacDonald, and J.P. Wikswo, “A Low Temperature Transfer of ALH84001 from Mars to Earth,” Science 290:791-895, 2000. 125. J.L. Kirschvink, E.J. Gaidos, L.E. Bertani, N.J. Beukes, J. Gutzmer, L.N. Maepa, and R.E. Steinberger, “Paleoproterozoic Snowball Earth: Extreme Climatic and Geochemical Global Change and Its Biological Consequences,” Proc. Natl. Acad. Sci. USA 97:1400-1405, 2000. 126. A.J. Alvarez, M. Khanna, G.A. Toranzos, and G. Stotzky, “Amplification of DNA Bound on Clay Minerals,” Mol. Ecol. 7:775-778, 1998. 127. D.C. White, W.M. Davis, J.S. Nickels, J.D. King, and R.J. Bobbie, “Determination of the Sedimentary Microbial Biomass by Extractable Lipid Phosphate,” Oecologia 40:51-62, 1979.
OCR for page 145
Signs of Life: A Report Based on the April 2000 Workshop on Life Detection Techniques 128. D.L. Balkwill, F.R. Leach, J.T. Wilson, J.F. McNabb, and D.C. White, “Equivalence of Microbial Biomass Measures Based on Membrane Lipid and Cell Wall Components, Adenosine Triphosphate, and Direct Counts in Subsurface Sediments,” Microbial Ecology 16:73-84, 1988. 129. M.M. Moeseneder, J.M. Arrieta, G. Muyzer, C. Winter, and G.J. Herndl, “Optimization of Terminal-restriction Fragment Length Polymorphism Analysis for Complex Marine Bacterioplankton Communities and Comparison with Denaturing Gradient Gel Electrophoresis,” Appl. Environ. Microbiol. 65:3518-3525, 1999. 130. G.E.C. Sheridan, C.I. Masters, J.A. Shallcross, and B.M. Mackey, “Detection of mRNA by Reverse Transcription PCR as an Indicator of Viability in Escherichia coli Cells,” Appl. Environ. Microbiol. 64(4):1313-1318, 1998. 131. C.A. Davidson, C.J. Griffith, A.C. Peters, and L.M. Fielding, “Evaluation of Two Methods for Monitoring Surface Cleanliness—ATP Bioluminescence and Traditional Hygiene Swabbing,” Luminescence 14(1):33-38, 1999. 132. V.G. Frundzhyan, L.Y. Brovko, V.S. Babunova, V.M. Kartashova, and N.N. Ugarova, “A Bioluminescence Assay of Total Bacterial Contamination of Fresh Milk,” Appl. Biochem. Microbiol. 35:321-327, 1999. 133. D.E. Atkinson, “The Energy Charge of the Adenylate Pool as a Regulatory Parameter. Interactions with Feedback Modifiers,” Biochemistry 7:4030-4034, 1968. 134. D.C. White, W.M. Davis, J.S. Nickels, J.D. King, and R.J. Bobbie, “Determination of the Sedimentary Microbial Biomass by Extractable Lipid Phosphate,” Oecologia 40:51-62, 1979. 135. D.L. Balkwill, F.R. Leach, J.T. Wilson, J.F. McNabb, and D.C. White, “Equivalence of Microbial Biomass Measures Based on Membrane Lipid and Cell Wall Components, Adenosine Triphosphate, and Direct Counts in Subsurface Sediments,” Microbial Ecology 16:73-84, 1988. 136. J.P. Diaper and C. Edwards, “The Use of Fluorogenic Esters to Detect Viable Bacteria by Flow-cytometry, ” Journal of Applied Bacteriology 77(2):221-228, 1994. 137. K. Venkateswaran, personal communication. 138. E. Zuckerkandl and L. Pauling, “Molecules as Documents of Evolutionary History,” Journal of Theoretical Biology 8:357-366, 1965. 139. S. Yamamoto and S. Harayama, “PCR Amplification and Direct Sequencing of Gyrb Genes with Universal Primers and Their Application to the Detection and Taxonomic Analysis of Pseudomonas putida Strains,” Appl. Environ. Microbiol. 61(3):1104-1109, 1995. 140. K. Venkateswaran, D.P. Moser, M.E. Dollhopf, D.P. Lies, D.A. Saffarini, B.J. MacGregor, D.B. Ringelberg, D.C. White, M. Nishijima, H. Sano, J. Burghardt, E. Stackebrandt, and K.H. Nealson, “Polyphasic Taxonomy of the Genus Shewanella and Description of Shewanella oneidensis sp. nov.,” International Journal of Systematic Bacteriology 49:705-724, 1999. 141. D. Jones and N.R. Krieg, “Serology and Chemotaxonomy,” in Bergey's Manual of Systematic Bacteriology, N.R. Krieg and J.G. Holt (eds.), Vol. 1, Williams & Wilkins, Baltimore, 1984, pp. 15-18. 142. M. Goodfellow and A.G. O'Donnell, “Roots of Bacterial Systematics,” in Handbook of New Bacterial Systematics, M. Goodfellow and A.G. O'Donnell (eds.), Academic Press, London, 1993, pp. 3-54. 143. C.R. Woese, “Bacterial Evolution,” Microbiol. Rev. 51:221-271, 1987. 144. N.R. Pace, D.A. Stahl, D.J. Lane, and G.J. Olsen, “The Analysis of Natural Microbial Populations by Ribosomal RNA Sequences, ” Advances in Microbial Ecology 9:1-55, 1986. 145. D.A. Stahl, “Evolution, Ecology and Diagnosis: Unity in Variety,” Bio/Technology 4:623-628, 1986. 146. D.A. Stahl and R. Amann, “Development and Application of Nucleic Acid Probes in Bacterial Systematics, ” in Sequencing and Hybridization Techniques in Bacterial Systematics, E. Stackebrandt and M. Goodfellow (eds.), John Wiley and Sons, Chichester, England, 1991, pp. 205-248. 147. D.M. Ward, M.M. Bateson, R. Weller, and A.L. Ruff-Roberts, “Ribosomal RNA Analysis of Microorganisms as They Occur in Nature, ” Advances in Microbial Ecology 12:219-286, 1992. 148. P. Hugenholtz, B.M. Goebel, and N.R. Pace, “Impact of Culture-independent Studies on the Emerging Phylogenetic View of Bacterial Diversity,” Journal Bacteriol. 180:4765-4774, 1988. 149. C.R. Woese, “Bacterial Evolution,” Microbiol. Rev. 51:221-271, 1987. 150. C.R. Woese, “Bacterial Evolution,” Microbiol. Rev. 51:221-271, 1987. 151. D.A. Stahl, “Application of Phylogenetically Based Hybridization Probes to Microbial Ecology,” Mol. Ecol. 4:535-542, 1995. 152. See <http://rdp.cme.msu.edu>. 153. B.L. Maidak, J.R. Cole, T.G. Lilburn, C.T. Parker, P.R. Saxman, J.M. Stredwick, G.M. Garrity, B. Li, G.J. Olsen, S. Pramanik, T.M. Schmidt, and J.M. Tiedje, “The RDP (Ribosomal Database Project) Continues,” Nucleic Acids Research 28(1):173-174, 2000. 154. B.L. Maidak, J.R. Cole, C.T. Parker, G.M. Garrity, N. Larsen, B. Li, T.G. Lilburn, M.J. McCaughey, G.J. Olsen, R. Overbeek, S. Pramanik, T.M. Schmidt, J.M. Tiedje, and C.R. Woese, “A New Version of the RDP (Ribosomal Database Project),” Nucleic Acids Research 27(1):171-173, 1999. 155. B.L. Maidak, J.R. Cole, T.G. Lilburn, C.T. Parker, P.R. Saxman, J.M. Stredwick, G.M. Garrity, B. Li, G.J. Olsen, S. Pramanik, T.M. Schmidt, and J.M. Tiedje, “The RDP (Ribosomal Database Project) Continues,” Nucleic Acids Research 28(1):173-174, 2000. 156. W.-T. Liu, T.L. Marsh, H. Cheng, and L.J. Forney, “Characterization of Microbial Diversity by Determining Terminal Restriction Fragment Length Polymorphisms of 16S Ribosomal DNA,” Appl. Environ. Microbiol. 63:4516-4522, 1997. 157. W.-T. Liu, T.L. Marsh, H. Cheng, and L.J. Forney, “Characterization of Microbial Diversity by Determining Terminal Restriction Fragment Length Polymorphisms of 16S Ribosomal DNA,” Appl. Environ. Microbiol. 63:4516-4522, 1997.
OCR for page 146
Signs of Life: A Report Based on the April 2000 Workshop on Life Detection Techniques 158. M.M. Moeseneder, J.M. Arrieta, G. Muyzer, C. Winter, and G.J. Herndl, “Optimization of Terminal-restriction Fragment Length Polymorphism Analysis for Complex Marine Bacterioplankton Communities and Comparison with Denaturing Gradient Gel Electrophoresis,” Appl. Environ. Microbiol. 65:3518-3525, 1999. 159. D.A. Stahl and R. Amann, “Development and Application of Nucleic Acid Probes in Bacterial Systematics, ” in Sequencing and Hybridization Techniques in Bacterial Systematics, E. Stackebrandt and M. Goodfellow (eds.), John Wiley and Sons, Chichester, U.K., 1991, pp. 205-248. 160. R.I. Amann, W. Ludwig, and K.-H. Schleifer, “Phylogenetic Identification and In Situ Detection of Individual Microbial Cells Without Cultivation,” Microbiol. Rev. 59:143-169, 1995. 161. E.F. DeLong, G.S. Wickham, and N.R. Pace, “Phylogenetic Stains: Ribosomal RNA-based Probes for the Identification of Single Cells,” Science 243:1360-1363, 1989. 162. R.I. Amann, W. Ludwig, and K.-H. Schleifer, “Phylogenetic Identification and In Situ Detection of Individual Microbial Cells Without Cultivation,” Microbiol. Rev. 59:143-169, 1995. 163. D.A. Stahl, “Application of Phylogenetically Based Hybridization Probes to Microbial Ecology,” Mol. Ecol. 4:535-542, 1995. 164. D.A. Stahl and R. Amann, “Development and Application of Nucleic Acid Probes in Bacterial Systematics, ” in Sequencing and Hybridization Techniques in Bacterial Systematics, E. Stackebrandt and M. Goodfellow (eds.), John Wiley and Sons, Chichester, England, 1991, pp. 205-248. 165. H. Hennecke, K. Kaluza, B. Thony, M. Fuhrmann, W. Ludwig, and E. Stackebrandt, “Concurrent Evolution of Nitrogenase Genes and 16S rRNA in Rhizobium Species and Other Nitrogen-fixing Bacteria,” Arch. Microbiol. 142:342-348, 1985. 166. C. Wawer and G. Muyzer, “Genetic Diversity of Desulfovibrio spp. in Environmental Samples Analyzed by Denaturing Gradient Gel Electrophoresis of [NiFe] Hydrogenase Gene Fragments,” Appl. Environ. Microbiol. 61:2203-2210, 1995. 167. M. Wagner, A.J. Roger, J.L. Flax, G.A. Brusseau, and D.A. Stahl, “Phylogeny of Dissimilatory Sulfite Reductases Supports an Early Origin of Sulfate Respiration,” Journal Bacteriol. 180:2975-2982, 1998. 168. D. Minz, J.L. Flax, S.J. Green, G. Muyzer, Y. Cohen, M. Wagner, B.E. Rittmann, and D.A. Stahl, “Diversity of Sulfate-reducing Bacteria in Oxic and Anoxic Regions of a Microbial Mat Characterized by Comparative Analysis of Dissimilatory Sulfite Reductase Genes,” Appl. Environ. Microbiol. 65:4666-4671, 1999. 169. D.M. Karl, D.F. Bird, K. Bjorkman, T. Houlihan, R. Shackelford, and L. Tupas, “Microorganisms in the Accreted Ice of Lake Vostok, Antarctica,” Science 286:2144-2147, 1999. 170. N.R. Pace, B.C. Thomas, and C.R. Woese, “Probing RNA Structure, Function, and History by Comparative Analysis, ” in The RNA World: The Nature of Modern RNA Suggests a Prebiotic RNA, R.F. Gesteland, T.R. Cech, and J.F. Atkins (eds.), 2nd ed., Cold Spring Harbor Laboratory Press, Cold Spring Harbor, New York, 1999, pp. 79-112. 171. N.R. Pace, “Origin of Life—Facing Up to the Physical Setting,” Cell 65:531-533, 1991. 172. S.M. Barns, R.E. Fundyga, M.W. Jeffries, and N.R. Pace, “Remarkable Archaeal Diversity Detected in a Yellowstone National Park Hot Spring Environment,” Proc. Natl. Acad. Sci. USA 91:1609-1613, 1994. 173. S.M. Barns, R.E. Fundyga, M.W. Jeffries, and N.R. Pace, “Remarkable Archaeal Diversity Detected in a Yellowstone National Park Hot Spring Environment,” Proc. Natl. Acad. Sci. USA 91:1609-1613, 1994. 174. W.L. Nicholson, N. Munakata, G. Horneck, H.J. Melosh, and P. Setlow, “Resistance of Bacillus Endospores to Extreme Terrestrial and Extraterrestrial Environments,” Microbiol. Mol. Biol. Rev. 64:548-572, 2000. 175. C. Mileikowsky, F.A. Cucinotta, J.W. Wilson, B. Gladman, G. Horneck, L. Lindegren, J. Melosh, H. Rickman, M. Valtonen, and J.Q. Zheng, “Natural Transfer of Viable Microbes in Space. 1. From Mars to Earth and Earth to Mars,” Icarus 145:391-427, 2000. 176. J.W. Valley, J.M. Eiler, C.M. Graham, E.K. Gibson, C.S. Romanek, and E.M. Stolper, “Low-temperature Carbonate Concretions in the Martian Meteorite ALH84001: Evidence from Stable Isotopes and Mineralogy,” Science 275:1633-1638, 1997. 177. E.R. Scott, A. Yamaguchi, and A.N. Krot, “Petrological Evidence for Shock Melting of Carbonates in the Martian Meteorite ALH84001,” Nature 387:377-379, 1997. 178. S.J. Mojzsis, R. Krishnamurthy, and G. Arrhenius, “Before RNA and After: Geophysical and Geochemical Constraints on Molecular Evolution,” in The RNA World: The Nature of Modern RNA Suggests a Prebiotic RNA , R.F. Gesteland, T.R. Cech, and J.F. Atkins (eds.), 2nd ed., Cold Spring Harbor Laboratory Press, Cold Spring Harbor, New York, 1999, pp. 1-48. 179. K.S. Makarova, L. Aravind, M.Y. Galperin, N.V. Grishin, R.L. Tatusov, Y.I. Wolf, and E.V. Koonin, “Comparative Genomics of the Archaea (Euryarchaeota): Evolution of Conserved Protein Families, the Stable Core, and the Variable Shell, ” Genome Research 9:608-628, 1999. 180. P.D. Ward and D. Brownlee, Rare Earth: Why Complex Life Is Uncommon in the Universe, Copernicus, New York, 2000. 181. W.L. Nicholson, N. Munakata, G. Horneck, H.J. Melosh, and P. Setlow, “Resistance of Bacillus Endospores to Extreme Terrestrial and Extraterrestrial Environments,” Microbiol. Mol. Biol. Rev. 64:548-572, 2000. 182. K. Pedersen, “Exploration of Deep Intraterrestrial Microbial Life: Current Perspectives, ” FEMS Microbiol. Lett. 185:9-16, 2000. 183. M.C. Malin and K.S. Edgett, “Evidence for Recent Groundwater Seepage and Surface Runoff on Mars, ” Science 288:2330-2335, 2000. 184. M.C. Malin, A. McEwen, M. Carr, L. Soderblom, P. Thomas, E. Danielson, P. James, J. Veverka, and W.K. Hartmann, “Evidence for Recent Volcanism on Mars from Crater Counts,” Nature 397:586-589, 1999.
Representative terms from entire chapter: