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4.5: Archaea - Biology

4.5: Archaea - Biology


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Learning Objectives

  • Describe the unique features of each category of Archaea
  • Explain why archaea might not be associated with human microbiomes or pathology
  • Give common examples of archaea commonly associated with unique environmental habitats

Like organisms in the domain Bacteria, organisms of the domain Archaea are all unicellular organisms. However, archaea differ structurally from bacteria in several significant ways, as discussed in Unique Characteristics of Prokaryotic Cells. To summarize:

  • The archaeal cell membrane is composed of ether linkages with branched isoprene chains (as opposed to the bacterial cell membrane, which has ester linkages with unbranched fatty acids).
  • Archaeal cell walls lack peptidoglycan, but some contain a structurally similar substance called pseudopeptidoglycan or pseudomurein.
  • The genomes of Archaea are larger and more complex than those of bacteria.

Domain Archaea is as diverse as domain Bacteria, and its representatives can be found in any habitat. Some archaea are mesophiles, and many are extremophiles, preferring extreme hot or cold, extreme salinity, or other conditions that are hostile to most other forms of life on earth. Their metabolism is adapted to the harsh environments, and they can perform methanogenesis, for example, which bacteria and eukaryotes cannot.

The size and complexity of the archaeal genome makes it difficult to classify. Most taxonomists agree that within the Archaea, there are currently five major phyla: Crenarchaeota, Euryarchaeota, Korarchaeota, Nanoarchaeota, and Thaumarchaeota. There are likely many other archaeal groups that have not yet been systematically studied and classified.

With few exceptions, archaea are not present in the human microbiota, and none are currently known to be associated with infectious diseases in humans, animals, plants, or microorganisms. However, many play important roles in the environment and may thus have an indirect impact on human health.

Crenarchaeota

Crenarchaeota is a class of Archaea that is extremely diverse, containing genera and species that differ vastly in their morphology and requirements for growth. All Crenarchaeota are aquatic organisms, and they are thought to be the most abundant microorganisms in the oceans. Most, but not all, Crenarchaeota are hyperthermophiles; some of them (notably, the genus Pyrolobus) are able to grow at temperatures up to 113 °C.1

Archaea of the genus Sulfolobus (Figure (PageIndex{1})) are thermophiles that prefer temperatures around 70–80°C and acidophiles that prefer a pH of 2–3.2 Sulfolobus can live in aerobic or anaerobic environments. In the presence of oxygen, Sulfolobus spp. use metabolic processes similar to those of heterotrophs. In anaerobic environments, they oxidize sulfur to produce sulfuric acid, which is stored in granules. Sulfolobus spp. are used in biotechnology for the production of thermostable and acid-resistant proteins called affitins.3 Affitins can bind and neutralize various antigens (molecules found in toxins or infectious agents that provoke an immune response from the body).

Another genus, Thermoproteus, is represented by strictly anaerobic organisms with an optimal growth temperature of 85 °C. They have flagella and, therefore, are motile. Thermoproteus has a cellular membrane in which lipids form a monolayer rather than a bilayer, which is typical for archaea. Its metabolism is autotrophic. To synthesize ATP, Thermoproteus spp. reduce sulfur or molecular hydrogen and use carbon dioxide or carbon monoxide as a source of carbon. Thermoproteus is thought to be the deepest-branching genus of Archaea, and thus is a living example of some of our planet’s earliest forms of life.

Exercise (PageIndex{1})

What types of environments do Crenarchaeota prefer?

Euryarchaeota

The phylum Euryarchaeota includes several distinct classes. Species in the classes Methanobacteria, Methanococci, and Methanomicrobia represent Archaea that can be generally described as methanogens. Methanogens are unique in that they can reduce carbon dioxide in the presence of hydrogen, producing methane. They can live in the most extreme environments and can reproduce at temperatures varying from below freezing to boiling. Methanogens have been found in hot springs as well as deep under ice in Greenland. Some scientists have even hypothesized that methanogens may inhabit the planet Mars because the mixture of gases produced by methanogens resembles the makeup of the Martian atmosphere.4

Methanogens are thought to contribute to the formation of anoxic sediments by producing hydrogen sulfide, making “marsh gas.” They also produce gases in ruminants and humans. Some genera of methanogens, notably Methanosarcina, can grow and produce methane in the presence of oxygen, although the vast majority are strict anaerobes.

The class Halobacteria (which was named before scientists recognized the distinction between Archaea and Bacteria) includes halophilic (“salt-loving”) archaea. Halobacteria require a very high concentrations of sodium chloride in their aquatic environment. The required concentration is close to saturation, at 36%; such environments include the Dead Sea as well as some salty lakes in Antarctica and south-central Asia. One remarkable feature of these organisms is that they perform photosynthesis using the protein bacteriorhodopsin, which gives them, and the bodies of water they inhabit, a beautiful purple color (Figure (PageIndex{2})).

Notable species of Halobacteria include Halobacterium salinarum, which may be the oldest living organism on earth; scientists have isolated its DNA from fossils that are 250 million years old.5 Another species, Haloferax volcanii, shows a very sophisticated system of ion exchange, which enables it to balance the concentration of salts at high temperatures

Exercise (PageIndex{2})

Where do Halobacteria live?

FINDING A LINK BETWEEN ARCHAEA AND DISEASE

Archaea are not known to cause any disease in humans, animals, plants, bacteria, or in other archaea. Although this makes sense for the extremophiles, not all archaea live in extreme environments. Many genera and species of Archaea are mesophiles, so they can live in human and animal microbiomes, although they rarely do. As we have learned, some methanogens exist in the human gastrointestinal tract. Yet we have no reliable evidence pointing to any archaean as the causative agent of any human disease.

Still, scientists have attempted to find links between human disease and archaea. For example, in 2004, Lepp et al. presented evidence that an archaean called Methanobrevibacter oralis inhabits the gums of patients with periodontal disease. The authors suggested that the activity of these methanogens causes the disease.6 However, it was subsequently shown that there was no causal relationship between M. oralis and periodontitis. It seems more likely that periodontal disease causes an enlargement of anaerobic regions in the mouth that are subsequently populated by M. oralis.7

There remains no good answer as to why archaea do not seem to be pathogenic, but scientists continue to speculate and hope to find the answer.

Summary

  • Archaea are unicellular, prokaryotic microorganisms that differ from bacteria in their genetics, biochemistry, and ecology.
  • Some archaea are extremophiles, living in environments with extremely high or low temperatures, or extreme salinity.
  • Only archaea are known to produce methane. Methane-producing archaea are called methanogens.
  • Halophilic archaea prefer a concentration of salt close to saturation and perform photosynthesis using bacteriorhodopsin.
  • Some archaea, based on fossil evidence, are among the oldest organisms on earth.
  • Archaea do not live in great numbers in human microbiomes and are not known to cause disease.
  1. E. Blochl et al.“Pyrolobus fumani, gen. and sp. nov., represents a novel group of Archaea, extending the upper temperature limit for life to 113°C.” Extremophiles 1 (1997):14–21.
  2. T.D. Brock et al. “Sulfolobus: A New Genus of Sulfur-Oxidizing Bacteria Living at Low pH and High Temperature.” Archiv für Mikrobiologie 84 no. 1 (1972):54–68.
  3. S. Pacheco et al. “Affinity Transfer to the Archaeal Extremophilic Sac7d Protein by Insertion of a CDR.” Protein Engineering Design and Selection 27 no. 10 (2014):431-438.
  4. R.R. Britt “Crater Critters: Where Mars Microbes Might Lurk.” www.space.com/1880-crater-cri...obes-lurk.html. Accessed April 7, 2015.
  5. H. Vreeland et al. “Fatty acid and DA Analyses of Permian Bacterium Isolated From Ancient Salt Crystals Reveal Differences With Their Modern Relatives.” Extremophiles 10 (2006):71–78.
  6. P.W. Lepp et al. “Methanogenic Archaea and Human Gum Disease.” Proceedings of the National Academies of Science of the United States of America 101 no. 16 (2004):6176–6181.
  7. R.I. Aminov. “Role of Archaea in Human Disease.” Frontiers in Cellular and Infection Microbiology 3 (2013):42.

Contributor

  • Nina Parker, (Shenandoah University), Mark Schneegurt (Wichita State University), Anh-Hue Thi Tu (Georgia Southwestern State University), Philip Lister (Central New Mexico Community College), and Brian M. Forster (Saint Joseph’s University) with many contributing authors. Original content via Openstax (CC BY 4.0; Access for free at https://openstax.org/books/microbiology/pages/1-introduction)


The desire to understand and distinguish the relative growth and activity of ammonia oxidising archaea (AOA) and ammonia oxidising bacteria (AOB) in soil nitrification has increased the search for selective inhibitors of these two groups. This study aimed to investigate the potency and specificity of simvastatin as a specific AOA inhibitor in pure cultures and in soil and to determine the effect of AOA inhibition on both ammonia oxidation activity and growth of AOB, under the hypothesis that AOB growth is higher when competition for NH4 + from AOA is removed. Simvastatin selectively inhibited pure cultures of all tested AOA at concentrations of 8–100 μM. In soil microcosms incubated for 21 days with low and high NH4 + concentrations, AOA but not AOB were selectively inhibited by simvastatin in both acidic (pH 4.5) and near-neutral (pH 6.5) soils. Additionally, growth of AOB significantly increased at both NH4 + concentrations following inhibition of AOA by simvastatin, suggesting that competition for substrate between AOA and AOB is a key factor restraining AOB growth in NH4 + -limited soils. Simvastatin can therefore be used as a selective AOA inhibitor to investigate kinetic characteristics of AOB in soils and to study competition between AOA and AOB in complex environments.

Present address: Institute for Food and Agricultural Sciences (IFAS), Department of Microbiology & Cell Science, University of Florida, 3205 College Avenue, Fort Lauderdale (Davie), FL33314, USA.

Present address: Department of Microbiology, Faculty of Science, Adekunle Ajasin University Akungba Akoko, Nigeria.


Proposal of the reverse flow model for the origin of the eukaryotic cell based on comparative analyses of Asgard archaeal metabolism

The origin of eukaryotes represents an unresolved puzzle in evolutionary biology. Current research suggests that eukaryotes evolved from a merger between a host of archaeal descent and an alphaproteobacterial endosymbiont. The discovery of the Asgard archaea, a proposed archaeal superphylum that includes Lokiarchaeota, Thorarchaeota, Odinarchaeota and Heimdallarchaeota suggested to comprise the closest archaeal relatives of eukaryotes, has helped to elucidate the identity of the putative archaeal host. Whereas Lokiarchaeota are assumed to employ a hydrogen-dependent metabolism, little is known about the metabolic potential of other members of the Asgard superphylum. We infer the central metabolic pathways of Asgard archaea using comparative genomics and phylogenetics to be able to refine current models for the origin of eukaryotes. Our analyses indicate that Thorarchaeota and Lokiarchaeota encode proteins necessary for carbon fixation via the Wood-Ljungdahl pathway and for obtaining reducing equivalents from organic substrates. By contrast, Heimdallarchaeum LC2 and LC3 genomes encode enzymes potentially enabling the oxidation of organic substrates using nitrate or oxygen as electron acceptors. The gene repertoire of Heimdallarchaeum AB125 and Odinarchaeum indicates that these organisms can ferment organic substrates and conserve energy by coupling ferredoxin reoxidation to respiratory proton reduction. Altogether, our genome analyses suggest that Asgard representatives are primarily organoheterotrophs with variable capacity for hydrogen consumption and production. On this basis, we propose the 'reverse flow model', an updated symbiogenetic model for the origin of eukaryotes that involves electron or hydrogen flow from an organoheterotrophic archaeal host to a bacterial symbiont.


4.5 Cytoskeleton

In this section, you will explore the following questions:

Connection for AP ® Courses

All cells, from simple bacteria to complex eukaryotes, possess a cytoskeleton composed of different types of protein elements, including microfilaments, intermediate filaments, and microtubules. The cytoskeleton serves a variety of purposes: provides rigidity and shape to the cell, facilitates cellular movement, anchors the nucleus and other organelles in place, moves vesicles through the cell, and pulls replicated chromosomes to the poles of a dividing cell. These protein elements are also integral to the movement of centrioles, flagella, and cilia.

The information presented and the examples highlighted in the section support concepts and Learning Objectives outlined in Big Idea 1 of the AP Biology Curriculum Framework, as shown in the table below.

The Learning Objectives listed in the Curriculum Framework provide a transparent foundation for the AP ® Biology course, an inquiry-based laboratory experience, instructional activities, and AP ® exam questions. A Learning Objective merges required content with one or more of the seven Science Practices.

Big Idea 1 The process of evolution drives the diversity and unity of life.
Enduring Understanding 1.B Organisms are linked by lines of descent from common ancestry
Essential Knowledge 1.B.1 Organisms share many conserved core processes and features that evolved and are widely distributed among organisms today.
Science Practice 7.2 The student can connect concepts in and across domain(s) to generalize or extrapolate in and/or across enduring understandings and/or big ideas.
Learning Objective 1.15 The student is able to describe specific examples of conserved core biological processes and features shared by all domains or within one domain of life and how these shared, conserved core processes and features support the concept of common ancestry for all organisms.

Teacher Support

Describe the cytoskeleton both as a “skeleton” because it provides the cell with shape and as “muscles” because it allows cells to move. The subunits of the cytoskeleton assemble and disassemble constantly, which is hard to imagine. Stress the concept of “dynamic equilibrium.” A vivid animation may illustrate the point better. Another way to demonstrate this is to have a few students stand in a line and then have a “pool” of students stand nearby. Then, on by one, ask students from the pool to exchange places with a student standing in the line. The line itself can grow and shrink by adding or taking away students in the line, respectively.

Both prokaryotic and eukaryotic cells possess cytoskeletons involved in cell division and shape maintenance. Although the molecular structures of cytoskeleton proteins are similar between two types of cells, the actual amino acid sequences of these proteins show very low levels of homology between the cytoskeleton proteins in prokaryotes and eukaryotes.

The two systems are not closely related.

Although the cytoskeleton of prokaryotes was discovered in the mid 90’s, the misconception that prokaryotes do not have a cytoskeleton is still widespread.

Microfilaments

If you were to remove all the organelles from a cell, would the plasma membrane and the cytoplasm be the only components left? No. Within the cytoplasm, there would still be ions and organic molecules, plus a network of protein fibers that help maintain the shape of the cell, secure some organelles in specific positions, allow cytoplasm and vesicles to move within the cell, and enable cells within multicellular organisms to move. Collectively, this network of protein fibers is known as the cytoskeleton . There are three types of fibers within the cytoskeleton: microfilaments, intermediate filaments, and microtubules (Figure 4.22). Here, we will examine each.

Of the three types of protein fibers in the cytoskeleton, microfilaments are the narrowest. They function in cellular movement, have a diameter of about 7 nm, and are made of two intertwined strands of a globular protein called actin (Figure 4.23). For this reason, microfilaments are also known as actin filaments.

Actin is powered by ATP to assemble its filamentous form, which serves as a track for the movement of a motor protein called myosin. This enables actin to engage in cellular events requiring motion, such as cell division in eukaryotic cells and cytoplasmic streaming, which is the circular movement of the cell cytoplasm in plant cells. Actin and myosin are plentiful in muscle cells. When your actin and myosin filaments slide past each other, your muscles contract.

Microfilaments also provide some rigidity and shape to the cell. They can depolymerize (disassemble) and reform quickly, thus enabling a cell to change its shape and move. White blood cells (your body’s infection-fighting cells) make good use of this ability. They can move to the site of an infection and phagocytize the pathogen.

Link to Learning

To see an example of a white blood cell in action, click here and watch a short time-lapse video of the cell capturing two bacteria. It engulfs one and then moves on to the other.

  1. The body’s immune system would not be affected by this.
  2. The body’s immune system would not be able to fight off pathogens like bacteria with fewer white blood cells. This can increase the risk of illness in HIV patients.
  3. The body’s immune system, in order to recoup this loss, will produce more WBC’s.
  4. The body’s immune system will fight the pathogens more vigorously in order to compensate for the fewer white blood cells.

Intermediate Filaments

Intermediate filaments are made of several strands of fibrous proteins that are wound together (Figure 4.24). These elements of the cytoskeleton get their name from the fact that their diameter, 8 to 10 nm, is between those of microfilaments and microtubules.

Intermediate filaments have no role in cell movement. Their function is purely structural. They bear tension, thus maintaining the shape of the cell, and anchor the nucleus and other organelles in place. Figure 4.22 shows how intermediate filaments create a supportive scaffolding inside the cell.

The intermediate filaments are the most diverse group of cytoskeletal elements. Several types of fibrous proteins are found in the intermediate filaments. You are probably most familiar with keratin, the fibrous protein that strengthens your hair, nails, and the epidermis of the skin.

Microtubules

As their name implies, microtubules are small hollow tubes. The walls of the microtubule are made of polymerized dimers of α-tubulin and β-tubulin, two globular proteins (Figure 4.25). With a diameter of about 25 nm, microtubules are the widest components of the cytoskeleton. They help the cell resist compression, provide a track along which vesicles move through the cell, and pull replicated chromosomes to opposite ends of a dividing cell. Like microfilaments, microtubules can disassemble and reform quickly.

Microtubules are also the structural elements of flagella, cilia, and centrioles (the latter are the two perpendicular bodies of the centrosome). In fact, in animal cells, the centrosome is the microtubule-organizing center. In eukaryotic cells, flagella and cilia are quite different structurally from their counterparts in prokaryotes, as discussed below.

Flagella and Cilia

To refresh your memory, flagella (singular = flagellum) are long, hair-like structures that extend from the plasma membrane and are used to move an entire cell (for example, sperm, Euglena). When present, the cell has just one flagellum or a few flagella. When cilia (singular = cilium) are present, however, many of them extend along the entire surface of the plasma membrane. They are short, hair-like structures that are used to move entire cells (such as paramecia) or substances along the outer surface of the cell (for example, the cilia of cells lining the Fallopian tubes that move the ovum toward the uterus, or cilia lining the cells of the respiratory tract that trap particulate matter and move it toward your nostrils.)

Despite their differences in length and number, flagella and cilia share a common structural arrangement of microtubules called a “9 + 2 array.” This is an appropriate name because a single flagellum or cilium is made of a ring of nine microtubule doublets, surrounding a single microtubule doublet in the center (Figure 4.26).

Science Practice Connection for AP® Courses

Think About It

The ribosomes in bacterial cells and in human cells are made up of proteins and ribosomal RNA, suggesting that both kinds of cells share a common ancestor cell type. What are examples of other features of cells that provide evidence for common ancestry?

Teacher Support

This question is an application of Learning Objectives 1.15 and Science Practice 7.2 because shared conserved core biological processes and features support the concept of common ancestry for all organisms on Earth.

In both prokaryotes and eukaryotes, DNA is the hereditary material and has the same structure.

The cytosol is made of an aqueous gel. The membrane is a fluid bilayer in which proteins are embedded. Do not use the flagellum as an example. It is convergent evolution.

You have now completed a broad survey of the components of prokaryotic and eukaryotic cells. For a summary of cellular components in prokaryotic and eukaryotic cells, see Table 4.1.

Cell Component Function Present in Prokaryotes? Present in Animal Cells? Present in Plant Cells?
Plasma membrane Separates cell from external environment controls passage of organic molecules, ions, water, oxygen, and wastes into and out of cell Yes Yes Yes
Cytoplasm Provides turgor pressure to plant cells as fluid inside the central vacuole site of many metabolic reactions medium in which organelles are found Yes Yes Yes
Nucleolus Darkened area within the nucleus where ribosomal subunits are synthesized. No Yes Yes
Nucleus Cell organelle that houses DNA and directs synthesis of ribosomes and proteins No Yes Yes
Ribosomes Protein synthesis Yes Yes Yes
Mitochondria ATP production/cellular respiration No Yes Yes
Peroxisomes Oxidizes and thus breaks down fatty acids and amino acids, and detoxifies poisons No Yes Yes
Vesicles and vacuoles Storage and transport digestive function in plant cells No Yes Yes
Centrosome Unspecified role in cell division in animal cells source of microtubules in animal cells No Yes No
Lysosomes Digestion of macromolecules recycling of worn-out organelles No Yes No
Cell wall Protection, structural support and maintenance of cell shape Yes, primarily peptidoglycan No Yes, primarily cellulose
Chloroplasts Photosynthesis No No Yes
Endoplasmic reticulum Modifies proteins and synthesizes lipids No Yes Yes
Golgi apparatus Modifies, sorts, tags, packages, and distributes lipids and proteins No Yes Yes
Cytoskeleton Maintains cell’s shape, secures organelles in specific positions, allows cytoplasm and vesicles to move within cell, and enables unicellular organisms to move independently Yes Yes Yes
Flagella Cellular locomotion Some Some No, except for some plant sperm cells.
Cilia Cellular locomotion, movement of particles along extracellular surface of plasma membrane, and filtration Some Some No

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    Methods

    Sampling, DNA extraction, quality control for shotgun sequencing

    Samples were taken from the sulfidic spring called Mühlbacher Schwefelquelle, Isling (MSI), near Regensburg (coordinates: 48.98 N, 12.13 E) at a depth of 1 m through a time period of four months 11 . A two-side opened Schott flask allowing a continuous flow through of spring water (approximately 5,500 l h −1 ) was used to filter biofilm droplets with a polyethylene net 11 . Samples were transported to the laboratory on ice, concentrated via centrifugation (10 min, 13,800 × g) and stored at −80 °C. Metagenomic DNA was extracted using MoBio Power Biofilm DNA kit (MoBio, Carlsbad, USA). Quality with regard to length of metagenomic DNA was verified using agarose gel electrophoresis and the quantity of archaea and bacteria in the samples was determined via qPCR according to established protocols 11 . In brief, 16S rRNA gene copy numbers in 1 μl of metagenomic DNA are amplified using universal and domain-biased primers. For metagenomic library construction, DNA was subjected to paired-end sequencing at LGC Genomics, Berlin (Germany). Using two independent sequencing platforms, Illumina HiSeq 2000 and Roche 454 FLX Titanium, 426,869,764 (average length: 100 bases, approximate insert size: 300 bases) and 246,234 reads (average length 342 bases, approximate insert size: 2750±500 bases) were generated, respectively (LGC Genomics, Berlin, Germany). For the Illumina reads LGC Genomics carried out clipping of sequencing adapters, quality filtering (removal of reads containing Ns, trimming of reads at 3′-end to get a minimum average Phred quality score of 10 over a window of 10 bases and discarding reads shorter than 5 nt using CASAVA tool 1.8.2 http://support.illumina.com/sequencing/sequencing_software/casava.ilmn) and filtering for paired reads. A subsequent quality check of the LGC Genomics filtered Illumina reads using FASTQC (http://www.bioinformatics.babraham.ac.uk/projects/fastqc) showed that no additional filtering was necessary. For 454 reads the program sff_extract.py (http://bioinf.comav.upv.es/sff_extract) was used to convert the raw SFF-files to fastq format and clip ends with low quality or adaptor sequences.

    Reconstruction of 16S rRNA genes and abundance estimation in MSI samples

    This was performed as described in the Supplementary Methods.

    Binning and completeness estimation of genomes from MSI site

    The filtered Illumina paired-end reads were digitally normalized to a maximal k-mer coverage of 130 and a minimal k-mer coverage of 3, resulting in a total of 54,853,038 paired reads 47 . A 10% subset of the normalized reads together with the entire set of 454 paired-end reads was used for a hybrid assembly with MIRA 48 (for details, please see Supplementary Table 1 a comparison with other assembly strategies is summarized in Supplementary Table 4). Resulting contigs were filtered by a minimal average coverage cutoff of 10 and a minimal length cutoff of 3 kpbs. These quality cutoffs have been shown to be very conservative in order to avoid chimeric sequences in datasets 49 . In addition, the sequenced reads were mapped back to the SM1-MSI contigs. Filtered contigs were manually inspected for large deviations in the coverage distribution that might indicate chimeras. 22 contigs showed large deviations and were checked for proper read-pairing using Tablet (http://ics.hutton.ac.uk/tablet). Read-pairing in these contigs did not support chimera assumption, therefore, the contigs were retained. All contigs were searched against a custom set of marker genes from the eggNOG database 50 and blast 51 results were phylogenetically analyzed using MEGAN4 (ref. 52) (best hit, bit score cutoff 150). Contigs classified as archaeal were extracted and archaeal bins were further constrained based on GC-coverage plots 53 and phylogenetic placement of the marker genes, i.e. 16S rRNA and hamus subunit genes, which were identified using blastn 51 . These bins were used as custom training sets for Phymm and the PhymmBL algorithm was used for taxonomic classification of the complete assembly 54 . This re-classification of the metagenome expanded the bins by recruiting further contigs based on nucleotide composition. Preliminary annotation of genomes and completeness estimation are described in the Supplementary Methods.

    Annotation of SM1-MSI genome

    The SM1-MSI bin was re-annotated using the synteny-supported annotation platform MaGe 55,56 . Specific tools that supported the manual annotation and curation are provided in the Supplementary Methods.

    Sampling, assembly, annotation, binning of CG data

    Crystal Geyser is located on the east bank of the Green River, 6 km south of the town of Green River, Utah, USA (38° 56.3' N, 110° 8.1' W). 65 L of geyser water was collected as it erupted on November 6th and 8th, 2009. Water samples for metagenomics were filtered sequentially through 3.0 and 0.2 μm polyethersulfone filters (Pall Corporation, NY, USA) via a peristaltic pump, and filters were immediately frozen on dry ice in the field and then stored at −80°C in the laboratory until processing. DNA was extracted via the PowerMax Soil DNA Isolation Kit (MO BIO Laboratories, Inc., Carlsbad, CA). For sequencing, separate 500-bps insert-size libraries were constructed from DNA extracted from the 3.0 and 0.2 μm samples. For the analyses described here, we used sequencing information from nine 3 μm filters, labeled A through J (analysis of the 0.2 μm data and other aspects of the community composition will be reported elsewhere). These filters were essentially replicates, collected from subsamples of the water from the two eruptions.

    Libraries were sequenced on the Illumina HiSeq platform and resulted in 36 Gb of paired-end sequence (239,247,978 reads at 150 bp per read). Reads were trimmed for quality (using Sickle Sickle: A sliding-window, adaptive, quality-based trimming tool for FastQ files (Version 1.21), available at https://github.com/najoshi/sickle) and assembled using the IDBA_UD assembly software 57 using default parameters. Genes were predicted with Prodigal 58 on all contigs >5 kb, using the ‘meta’ option. Predicted proteins on each contig were assigned a preliminary functional annotation based on best blast hits to the UniRef90 database 59 . Because the SM1-CG genome was substantially more highly sampled than any other genome in the 3 μm samples, binning of the genome could be established using coverage and GC content. Multiple assemblies of data subsets were carried out to attempt to improve the genome quality by lowering the coverage, but this conferred no advantage. The best assembled genome came from sample I, and this was used for comparative genomic analysis with SM1-MSI genome. Here, blastp 51 was used to search for protein similarities between the genomes (e-value=10 −5 ) and annotations were only considered if both analyses pathways (for SM1-MSI and SM1-CG) produced identical functional predictions.

    We compared the representation of SM1-CG in samples A through I by evaluating coverage of scaffolds carrying ribosomal protein S3 (rpS3) genes (the three most abundant organisms in each sample were identical and at comparable relative coverage levels). The rpS3 sequences of SM1 from the samples were identical so only sequences from samples A, B and C were included in our phylogenomic analyses.

    The SM1-MSI genome (scaffold and protein sequences) is available as Supplementary Data 1 and 2.

    Phylogenomic analysis and estimation of horizontal gene transfer

    These are described in the Supplementary Methods.

    Transcriptomics of certain metabolic pathways in SM1-MSI

    For genes for certain key enzymes or for single subunits thereof, transcription was tested via specific mRNA detection in biofilm samples (Supplementary Table 5 containing list of genes and primers). Total RNA was isolated using the PowerBiofilm RNA Isolation Kit (Mobio Laboratories Inc., Carlsbad, USA) according to manufacturers’ instructions (DNA digestion was performed for 30 min). After precipitation of nucleic acids DNAse treatment was repeated, followed by subsequent reverse-transcription to cDNA (QuantiTect Rev. Transcription Kit, Qiagen, Hilden, Germany). Specific primers were designed using the web tool Primr3v.0.4.0 (http://biotools.umassmed.edu/bioapps/primer3_www.cgi parameters: product size: optimum 400 bp, GC% 40–60%, annealing temperature: 60 °C optimum). Specificity of primers was tested using blast 51 against NCBI NR and the metagenome. CDNA was used for amplification with designed primer pairs individually (denaturation time: 5 min 95 °C 30 cycles: 45 s 94 °C, 45 s 60 °C, 90 s 72 °C final elongation: 10 min 72 °C). Positive PCR products were purified (HiYield® Gel/PCR DNA Fragments Extraction Kit Süd-Laborbedarf GmbH, Gauting, Germany) and Sanger sequenced (LGC Genomics GmbH, Berlin, Germany). Experiments were carried out in duplicates.

    Fluorescence immuno-labeling of SM1-MSI cells in biofilms

    For the production of hamus-specific antibodies, hami filaments were released from the cells as follows. Biofilm samples were incubated in KPH buffer (NaCl 0.7 mM, MgCl2 0.1 mM, CaSO4 1.6 mM, HEPES 1.0 mM, supplemented with 0.1% SDS (v/v)) for 25 min and periodically vortexed, causing SM1 euryarchaeal cells to dissolve completely. The resulting suspension was centrifuged (30 min, 5,500 × g, 20 °C) to remove larger precipitates and the supernatant containing hami was ultracentrifuged (1 h, 92,387.1 × g, Beckman OPTIMA LE 80 K, 70.1 Ti-Rotor, 4 °C). The pellet was re-suspended in 2 ml of KPH buffer and applied on a sucrose-gradient (10–70% sucrose (w/v) in sterile KPH) and centrifuged for 17 h (309,000 × g, Beckman OPTIMA LE 80 K, SW 60 Rotor, 4 °C). The band appearing in the lower third of the tube was removed using a sterile syringe. After confirming the presence of hami via transmission electron microscopy (see below) the sample was sent to Davids biotechnology (Regensburg, Germany) for antibody production. A chicken was pre-immunized with the hami-fraction (0.22 mg/ml) three times over the course of 21 days. 28 days after the first immunization, eggs were collected and the IgG fraction (‘anti-hamus’ 15.1 mg/ml in 0.02% sodium-azide) was harvested.

    For immuno-staining, collected biofilms were fixed with paraformaldehyde (5% (v/v)) at room temperature (1 h) and washed three times with 1x PBS (phosphate buffered saline). Afterwards, fixed cells were incubated at 30 °C in PBST (PBS including Tween20 0.05% (v/v) with 0.1% SDS (v/v)) for 15 min, followed by a centrifugation step (15 min, 14,500 × g, 20 °C). The first primary antibody, anti-Anabaena FtsZ 60 (AS07217, Agrisera dilution 1:200) was added and incubated at 30 °C for 2 h. Cells were centrifuged, followed by incubation in PBST (+0.1% SDS (v/v)) for 15 min and another centrifugation step. After incubation for 1 h with the conjugated goat anti-rabbit IgG (dilution 1:200), the cells were washed twice with PBST (+0.1% SDS (v/v)), spread within a well of a gelatine-coated slide (P. Marienfeld KG, Lauda-Koenigshofen, Germany) and fixed via air-drying. After incubation with 16 μl of PBST (+0.1% SDS (v/v)) at 37 °C, the PBST buffer was replaced with 16 μl PBST buffer containing the anti-hamus IgG (dilution 1:2,000) and the cells were labeled at 37 °C for 1 h. Subsequently, the slide was washed 15 min in PBST (+0.1% SDS (v/v)), rinsed with H2O and air dried. The second antibody (goat anti-chicken, Cy3-labeled 0.64 mg/ml, dilution 1:500) was added and incubated at 37 °C for 1 h. After washing two times with PBST (+0.1% SDS (v/v)), the slide was rinsed with H2O, DAPI stained and analyzed using fluorescence microscopy (Olympus (BX53F, Hamburg, Germany) with epifluorescence equipment and imaging software cellSens).

    Transmission and scanning electron microscopy of MSI biofilms

    For TEM, fresh, unfixed biofilm pieces were deposited on a carbon-coated copper grid and negatively stained with 2% (w/v) uranyl acetate, pH 4.5 or 2.0% (w/v) phosphotungstic acid (PTA), pH 7.0. In a second approach, biofilms were treated with 1% SDS (w/v) for 30 min, causing destruction of the cell wall and thus a release of cell appendages. These samples were examined using a CM12 transmission electron microscope (FEI, Eindhoven, The Netherlands) operated at 120 keV. All images were digitally recorded using a slow-scan charge-coupled device camera that was connected to a computer with TVIPS software (TVIPS GmbH, Gauting, Germany).

    For conventional fixation, freshly taken biofilms were fixed in original spring water including 0.1% (w/v) glutardialdehyde. Samples were rinsed several times in fixative buffer and postfixed at room temperature for 1 h with 1% (w/v) osmium tetroxide. After two washing steps in water, the cells were stained for 30 min with 1% (w/v) uranyl acetate in 20% (v/v) acetone. Dehydration was performed by a graded acetone series. Samples were then infiltrated and embedded in Spurr’s low-viscosity resin. For high-pressure freezing experiment samples were frozen either with a Leica HPM100 (Leica Microsystems GmbH, Wetzlar, Germany, Fig. 5) or a Wohlwend HPF Compact 02 (Engineering Office M-Wohlwend GmbH, Sennwald, Switzerland Fig. 4). In the first case aluminum platelets were used which were filled with one piece of biofilm. Freeze substitution was performed in acetone with 2% (w/v) osmium tetroxide and 0.2% (w/v) uranyl acetate, including 5% (v/v) water. After embedding the samples in Spurr’s low-viscosity resin, ultrathin sections were cut with a diamond knife and mounted onto uncoated copper grids. The sections were poststained with aqueous lead citrate (100 mM, pH 13.0). Transmission electron micrographs of samples prepared this way were taken with an EM 912 electron microscope (Zeiss) equipped with an integrated OMEGA energy filter operated at 80 kV in the zero loss mode.

    For SEM, drops of the sample were placed onto a glass slide, covered with a coverslip, and rapidly frozen with liquid nitrogen. The coverslip was removed with a razor blade and the glass slide was immediately fixed with 2.5% (w/v) glutaraldehyde in 10 mM cacodylate buffer (pH 7.0), postfixed with 1% (w/v) osmium tetroxide in fixative buffer, dehydrated in a graded series of acetone solutions, and critical-point dried after transfer to liquid CO2. Specimens were mounted on stubs, coated with 3 nm platinum using a magnetron sputter coater, and examined with a Zeiss Auriga scanning electron microscope operated at 1–2 kV. When using the Wohlwend HPF Compact 02, biofilm aggregates were placed in the centre of 3 mm gold carriers and high pressure frozen with a and subsequently freeze substituted in an automatic EM AFS 2 unit (Leica Microsystems GmbH, Wetzlar, Germany). The substitution medium consisted of acetone in combination with 0.2% (w/v) osmium tetroxide, 0.25% (w/v) uranyl acetate and 5% (v/v) water. The substitution program including washing steps and the following epon embedding, thin sectioning and post staining was carried out as described previously 61 . For these samples, microscopy was performed on a JEOL JEM-2100 (JEOL, Tokyo, Japan) also operated at 120 kV and equipped with a 2 k × 2 k fast scan CCD camera F214 combined with EM Menu4 (TVIPS GmbH, Gauting, Gemany).

    FIB-SEM tomography of MSI biofilms

    The focused ion beam FIB block face serial sectioning was performed using a Zeiss-Auriga workstation. The focused ion beam consisted of Ga + ions accelerated by a voltage of 30 kV. In the cut-and-view mode, sections ranging in thickness between 5 nm and 10 nm (dependent on the magnification) were produced with the FIB and field emission scanning electron microscopy (FESEM) images, which were recorded at 1.5 kV using the in-lens energy selective backscattered (EsB) detector set to −1,200 V. Specimens were tilted to an angle of 54° images were tilt corrected for undistorted surface view.

    Lipid extraction and HPLC-MS analysis from biofilms

    Total lipid extracts (TLEs) were obtained from samples of the biofilm from MSI site using a modified Bligh and Dyer protocol 62 , after adding an internal standard (phosphatidylcholine C21:0/21:0) and 3 g of combusted sea sand. Approximately 10 8 to 10 9 cells were subjected to extraction. The obtained TLEs were stored at −20 °C and analysis of IPLs was performed by high-performance liquid chromatography electrospray ionization mass spectrometry (HPLC-ESI-MS). Separation of IPLs was achieved on a Dionex Ultimate 3000 UHPLC equipped with a Waters Acquity UPLC BEH Amide column (150 × 2.1 mm, 1.8 μm particle size). Chromatographic conditions, according to a previously published method 63 , were as follows: constant flow rate of 0.4 ml/min with eluent A (75% acetonitrile 25% DCM 0.01% formic acid 0.01% ammonium hydroxide solution (NH3aq)) and eluent B (50% MeOH 50% Milli-Q water 0.4% formic acid 0.4% NH3aq). Under a constant flow, the HPLC routine applied: 99% A and 1% B for 2.5 min, increasing to 5% B at 4 min, followed by a linear gradient to 25% B at 22.5 min and then to 40% B at 26.5 min. Thereafter a 1 min washing step with 40% B followed and afterwards reset to the initial conditions for 8 min to achieve column re-equilibration. Compound detection was conducted on a Bruker maXis Ultra-High Resolution qToF-MS, equipped with an ESI interface. IPLs were measured in positive ionization mode, while scanning a mass-to-charge (m/z) range of 150–2,000, with automated data-dependent MS/MS fragmentation of base peak ions. Compound identification was achieved by monitoring exact masses of possible parent ions (present mainly as H + and NH4 + adducts) in combination with characteristic fragmentation patterns 62,64 . The reported relative distribution of microbial lipids is based on the peak areas of the respective molecular ions without differentiating for potential differences in response factors the data should therefore be viewed as semi-quantitative.

    Stable carbon isotope analyses at MSI site

    Stable carbon isotopic (δ 13 C) composition of gases and lipids was obtained from groundwater samples and SM1 biofilm biomass. The δ 13 C values of CO2 and CH4 were measured from the headspace of groundwater samples (taken anaerobically and frozen on dry ice at MSI site by gas chromatography (GC) coupled to isotope-ratio mass spectrometry (irMS) (Trace GC ultra+DeltaPlus XP irMS, ThermoFinnigan). Given the dominance of glycolipids among both archaeal and bacterial lipids, TLEs were acid hydrolyzed (2.5% HCl in methanol) and derivatized with N,O-bis(trimethylsilyl)trifluoroacetamide in pyridine prior to GC-irMS (Trace GC Ultra coupled to a GC-IsoLink/ConFlow IV interface and a Delta V Plus irMS all from Thermo Scientific). δ 13 C values of lipids were corrected for additional carbon introduced during derivatization. The δ 13 C values are expressed versus Vienna PeeDee Belemnite (VPDB). Measurements of gases and lipids in GC-irMS were performed at least in duplicate and the analytical error was <0.5‰.

    NanoSIMS, Raman spectroscopy, and conductivity experiments

    These were performed on MSI biofilm samples as described in the Supplementary Methods.

    Analysis of diversity and distribution of the SM1 group

    This is presented in the Supplementary Methods, and all accession codes are provided in Supplementary Table 6.


    Contents

    Carl Woese was born in Syracuse, New York on July 15, 1928. Woese attended Deerfield Academy in Massachusetts. He received a bachelor's degree in mathematics and physics from Amherst College in 1950. During his time at Amherst, Woese took only one biology course (Biochemistry, in his senior year) and had "no scientific interest in plants and animals" until advised by William M. Fairbank, then an assistant professor of physics at Amherst, to pursue biophysics at Yale. [11]

    In 1953, he completed a Ph.D. in biophysics at Yale University, where his doctoral research focused on the inactivation of viruses by heat and ionizing radiation. [12] [13] He studied medicine at the University of Rochester for two years, quitting two days into a pediatrics rotation. [13] Then he became a postdoctoral researcher in biophysics at Yale University investigating bacterial spores. [14] From 1960–63, he worked as a biophysicist at the General Electric Research Laboratory in Schenectady, New York. [12] [15] In 1964, Woese joined the microbiology faculty of the University of Illinois at Urbana–Champaign, where he focused on Archaea, genomics, and molecular evolution as his areas of expertise. [10] [12] [15] He became a professor at the University of Illinois at Urbana–Champaign's Carl R. Woese Institute for Genomic Biology, which was renamed in his honor in 2015, after his death. [15]

    Woese died on December 30, 2012, following complications from pancreatic cancer, leaving as survivors his wife Gabriella and two sons. [16] [17] [18]

    Early work on the genetic code Edit

    Woese turned his attention to the genetic code while setting up his lab at General Electric's Knolls Laboratory in the fall of 1960. [13] Interest among physicists and molecular biologists had begun to coalesce around deciphering the correspondence between the twenty amino acids and the four letter alphabet of nucleic acid bases in the decade following James D. Watson, Francis Crick, and Rosalind Franklin's discovery of the structure of DNA in 1953. [19] Woese published a series of papers on the topic. In one, he deduced a correspondence table between what was then known as "soluble RNA" and DNA based upon their respective base pair ratios. [20] He then re-evaluated experimental data associated with the hypothesis that viruses used one base, rather than a triplet, to encode each amino acid, and suggested 18 codons, correctly predicting one for proline. [13] [21] Other work established the mechanistic basis of protein translation, but in Woese's view, largely overlooked the genetic code's evolutionary origins as an afterthought. [19]

    In 1962 Woese spent several months as a visiting researcher at the Pasteur Institute in Paris, a locus of intense activity on the molecular biology of gene expression and gene regulation. [13] While in Paris, he met Sol Spiegelman, who invited Woese to visit the University of Illinois after hearing his research goals at this visit Spiegelman offered Woese a position with immediate tenure beginning in the fall of 1964. [13] With the freedom to patiently pursue more speculative threads of inquiry outside the mainstream of biological research, Woese began to consider the genetic code in evolutionary terms, asking how the codon assignments and their translation into an amino acid sequence might have evolved. [13] [22]

    Discovery of the third domain Edit

    For much of the 20th century, prokaryotes were regarded as a single group of organisms and classified based on their biochemistry, morphology and metabolism. In a highly influential 1962 paper, Roger Stanier and C. B. van Niel first established the division of cellular organization into prokaryotes and eukaryotes, defining prokaryotes as those organisms lacking a cell nucleus. [23] [24] Adapted from Édouard Chatton's generalization, Stanier and Van Niel's concept was quickly accepted as the most important distinction among organisms yet they were nevertheless skeptical of microbiologists' attempts to construct a natural phylogenetic classification of bacteria. [25] However, it became generally assumed that all life shared a common prokaryotic (implied by the Greek root πρό (pro-), before, in front of) ancestor. [24] [26]

    In 1977, Carl Woese and George E. Fox experimentally disproved this universally held hypothesis about the basic structure of the tree of life. [27] Woese and Fox discovered a kind of microbial life which they called the “archaebacteria” (Archaea). [5] They reported that the archaebacteria comprised "a third kingdom" of life as distinct from bacteria as plants and animals. [5] Having defined Archaea as a new "urkingdom" (later domain) which were neither bacteria nor eukaryotes, Woese redrew the taxonomic tree. His three-domain system, based on phylogenetic relationships rather than obvious morphological similarities, divided life into 23 main divisions, incorporated within three domains: Bacteria, Archaea, and Eucarya. [3]

    Acceptance of the validity of Woese's phylogenetically valid classification was a slow process. Prominent biologists including Salvador Luria and Ernst Mayr objected to his division of the prokaryotes. [28] [29] Not all criticism of him was restricted to the scientific level. A decade of labor-intensive oligonucleotide cataloging left him with a reputation as "a crank," and Woese would go on to be dubbed as "Microbiology's Scarred Revolutionary" by a news article printed in the journal Science. [6] The growing body of supporting data led the scientific community to accept the Archaea by the mid-1980s. [13] Today, few scientists cling to the idea of a unified Prokarya.

    Woese's work on Archaea is also significant in its implications for the search for life on other planets. Before the discovery by Woese and Fox, scientists thought that Archaea were extreme organisms that evolved from the microorganisms more familiar to us. Now, most believe they are ancient, and may have robust evolutionary connections to the first organisms on Earth. [30] Organisms similar to those archaea that exist in extreme environments may have developed on other planets, some of which harbor conditions conducive to extremophile life. [31]

    Notably, Woese's elucidation of the tree of life shows the overwhelming diversity of microbial lineages: single-celled organisms represent the vast majority of the biosphere's genetic, metabolic, and ecologic niche diversity. [32] As microbes are crucial for many biogeochemical cycles and to the continued function of the biosphere, Woese's efforts to clarify the evolution and diversity of microbes provided an invaluable service to ecologists and conservationists. It was a major contribution to the theory of evolution and to our knowledge of the history of life. [19]

    Woese wrote, "My evolutionary concerns center on the bacteria and the archaea, whose evolutions cover most of the planet's 4.5-billion-year history. Using ribosomal RNA sequence as an evolutionary measure, my laboratory has reconstructed the phylogeny of both groups, and thereby provided a phylogenetically valid system of classification for prokaryotes. The discovery of the archaea was in fact a product of these studies". [12]

    Evolution of primary cell types Edit

    Woese also speculated about an era of rapid evolution in which considerable horizontal gene transfer occurred between organisms. [27] [33] First described by Woese and Fox in a 1977 paper and explored further with microbiologist Jane Gibson in a 1980 paper, these organisms, or progenotes, were imagined as protocells with very low complexity due to their error-prone translation apparatus ("noisy genetic transmission channel"), which produced high mutation rates that limited the specificity of cellular interaction and the size of the genome. [34] [35] This early translation apparatus would have produced a group of structurally similar, functionally equivalent proteins, rather than a single protein. [27] Furthermore, because of this reduced specificity, all cellular components were susceptible to horizontal gene transfer, and rapid evolution occurred at the level of the ecosystem. [33] [36]

    The transition to modern cells (the "Darwinian Threshold") occurred when organisms evolved translation mechanisms with modern levels of fidelity: improved performance allowed cellular organization to reach a level of complexity and connectedness that made genes from other organisms much less able to displace an individual's own genes. [33]

    In later years, Woese's work concentrated on genomic analysis to elucidate the significance of horizontal gene transfer (HGT) for evolution. [37] He worked on detailed analyses of the phylogenies of the aminoacyl-tRNA synthetases and on the effect of horizontal gene transfer on the distribution of those key enzymes among organisms. [38] The goal of the research was to explain how the primary cell types (the archaeal, eubacterial, and eukaryotic) evolved from an ancestral state in the RNA world. [12]

    Woese shared his thoughts on the past, present, and future of biology in Current Biology: [11]

    The "important questions" that 21st century biology faces all stem from a single question, the nature and generation of biological organization. . . . Yes, Darwin is back, but in the company of . . . scientists who can see much further into the depths of biology than was possible heretofore. It is no longer a "10,000 species of birds" view of evolution—evolution seen as a procession of forms. The concern is now with the process of evolution itself. [11]

    I see the question of biological organization taking two prominent directions today. The first is the evolution of (proteinaceous) cellular organization, which includes sub-questions such as the evolution of the translation apparatus and the genetic code, and the origin and nature of the hierarchies of control that fine-tune and precisely interrelate the panoply of cellular processes that constitute cells. It also includes the question of the number of different basic cell types that exist on earth today: did all modern cells come from a single ancestral cellular organization? [11]

    The second major direction involves the nature of the global ecosystem. . . . Bacteria are the major organisms on this planet—in numbers, in total mass, in importance to the global balances. Thus, it is microbial ecology that . . . is most in need of development, both in terms of facts needed to understand it, and in terms of the framework in which to interpret them. [11]

    Woese considered biology to have an "all-important" role in society. In his view, biology should serve a broader purpose than the pursuit of "an engineered environment": [11]

    What was formally recognized in physics needs now to be recognized in biology: science serves a dual function. On the one hand it is society's servant, attacking the applied problems posed by society. On the other hand, it functions as society's teacher, helping the latter to understand its world and itself. It is the latter function that is effectively missing today. [11]

    Woese was a MacArthur Fellow in 1984, was made a member of the National Academy of Sciences in 1988, received the Leeuwenhoek Medal (microbiology's highest honor) in 1992, the Selman A. Waksman Award in Microbiology in 1995 from the National Academy of Sciences, [39] and was a National Medal of Science recipient in 2000. In 2003, he received the Crafoord Prize from the Royal Swedish Academy of Sciences "for his discovery of a third domain of life". [40] [41] He was elected to the American Philosophical Society in 2004. [42] In 2006, he was made a foreign member of the Royal Society. [10]

    Many microbial species, such as Pyrococcus woesei, [43] Methanobrevibacter woesei, [44] and Conexibacter woesei, [45] are named in his honor.

    Microbiologist Justin Sonnenburg of Stanford University said "The 1977 paper is one of the most influential in microbiology and arguably, all of biology. It ranks with the works of Watson and Crick and Darwin, providing an evolutionary framework for the incredible diversity of the microbial world". [19]

    With regard to Woese's work on horizontal gene transfer as a primary evolutionary process, Professor Norman R. Pace of the University of Colorado at Boulder said, "I think Woese has done more for biology writ large than any biologist in history, including Darwin. There's a lot more to learn, and he's been interpreting the emerging story brilliantly". [46]


    Chapter Summary

    A cell is the smallest unit of life. Most cells are so tiny that they cannot be seen with the naked eye. Therefore, scientists use microscopes to study cells. Electron microscopes provide higher magnification, higher resolution, and more detail than light microscopes. The unified cell theory states that all organisms are composed of one or more cells, the cell is the basic unit of life, and new cells arise from existing cells.

    4.2 Prokaryotic Cells

    Prokaryotes are predominantly single-celled organisms of the domains Bacteria and Archaea. All prokaryotes have plasma membranes, cytoplasm, ribosomes, and DNA that is not membrane-bound. Most have peptidoglycan cell walls and many have polysaccharide capsules. Prokaryotic cells range in diameter from 0.1 to 5.0 μm.

    As a cell increases in size, its surface area-to-volume ratio decreases. If the cell grows too large, the plasma membrane will not have sufficient surface area to support the rate of diffusion required for the increased volume.

    4.3 Eukaryotic Cells

    Like a prokaryotic cell, a eukaryotic cell has a plasma membrane, cytoplasm, and ribosomes, but a eukaryotic cell is typically larger than a prokaryotic cell, has a true nucleus (meaning its DNA is surrounded by a membrane), and has other membrane-bound organelles that allow for compartmentalization of functions. The plasma membrane is a phospholipid bilayer embedded with proteins. The nucleus’s nucleolus is the site of ribosome assembly. Ribosomes are either found in the cytoplasm or attached to the cytoplasmic side of the plasma membrane or endoplasmic reticulum. They perform protein synthesis. Mitochondria participate in cellular respiration they are responsible for the majority of ATP produced in the cell. Peroxisomes hydrolyze fatty acids, amino acids, and some toxins. Vesicles and vacuoles are storage and transport compartments. In plant cells, vacuoles also help break down macromolecules.

    Animal cells also have a centrosome and lysosomes. The centrosome has two bodies perpendicular to each other, the centrioles, and has an unknown purpose in cell division. Lysosomes are the digestive organelles of animal cells.

    Plant cells and plant-like cells each have a cell wall, chloroplasts, and a central vacuole. The plant cell wall, whose primary component is cellulose, protects the cell, provides structural support, and gives shape to the cell. Photosynthesis takes place in chloroplasts. The central vacuole can expand without having to produce more cytoplasm.

    4.4 The Endomembrane System and Proteins

    The endomembrane system includes the nuclear envelope, lysosomes, vesicles, the ER, and Golgi apparatus, as well as the plasma membrane. These cellular components work together to modify, package, tag, and transport proteins and lipids that form the membranes.

    The RER modifies proteins and synthesizes phospholipids used in cell membranes. The SER synthesizes carbohydrates, lipids, and steroid hormones engages in the detoxification of medications and poisons and stores calcium ions. Sorting, tagging, packaging, and distribution of lipids and proteins take place in the Golgi apparatus. Lysosomes are created by the budding of the membranes of the RER and Golgi. Lysosomes digest macromolecules, recycle worn-out organelles, and destroy pathogens.

    4.5 The Cytoskeleton

    The cytoskeleton has three different types of protein elements. From narrowest to widest, they are the microfilaments (actin filaments), intermediate filaments, and microtubules. Microfilaments are often associated with myosin. They provide rigidity and shape to the cell and facilitate cellular movements. Intermediate filaments bear tension and anchor the nucleus and other organelles in place. Microtubules help the cell resist compression, serve as tracks for motor proteins that move vesicles through the cell, and pull replicated chromosomes to opposite ends of a dividing cell. They are also the structural element of centrioles, flagella, and cilia.

    4.6 Connections between Cells and Cellular Activities

    Animal cells communicate via their extracellular matrices and are connected to each other via tight junctions, desmosomes, and gap junctions. Plant cells are connected and communicate with each other via plasmodesmata.

    When protein receptors on the surface of the plasma membrane of an animal cell bind to a substance in the extracellular matrix, a chain of reactions begins that changes activities taking place within the cell. Plasmodesmata are channels between adjacent plant cells, while gap junctions are channels between adjacent animal cells. However, their structures are quite different. A tight junction is a watertight seal between two adjacent cells, while a desmosome acts like a spot weld.


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    78 Cards in this Set

    Despite the diversity of cell type and function, all cells have these three things in common:

    a) cytoplasm, DNA, and organellese

    b) a plasma membrane, DNA, and proteins

    c) cytoplasm, DNA, and a plasma membrane

    d) carbohydrates, nucleic acids, and proteins

    c) cytoplasm, DNA, and plasma membrane

    Cells differ in size, shape, and function, but all start out life with a plasma membrane, cytoplasm, and a region of DNA. Section 4.2

    Every cell is descended from another cell. This idea is part of __________.

    By the cell theory, all organisms consists of one or more cells the cell is the smallest unit of life each new cell arises from another, pre-existing cell and a cell passed hereditary material to it's offspring. Section 4.2

    The surface-to-volume ratio _________.

    a) does not apply to prokaryotic cells

    c) is part of the cell theory

    True or false? Ribosomes are only found in bacteria and archaea.

    Bacteria and archaea (prokaryotes) have ribosomes, and eukaryotic cells (animal and plant) have ribosomes in their endomembrane system. Section 4.4 and 4.7

    Unlike eukaryotic cells, bacterial cells __________.

    a) have no plasma membrane

    Bacteria and archaea, informally grouped as "prokaryotes," are the most diverse forms of life. These single-celled organisms have no nucleus, but they have nucleoids and ribosomes. Section 4.4.

    True or false? Some protists start out life with no nucleus.

    Protists are eukaryotes by definition, all eukaryotes start life with a nucleus. Section 4.5.

    Cell membranes consist mainly of a _________.

    a) a carbohydrate bilayer and proteins

    b) protein bilayer and phospholipids

    c) lipid bilayer and proteins

    c) lipid bilayer and proteins

    All cell membranes, including the plasma membrane and organelle members, are selectively permeable and consist mainly of phospholipids organized as a lipid bilayer. Many different proteins embedded in a bilayer or attached to one of its surfaces carry out membrane functions. Section 4.2.

    Enzymes contained in _________ break down worn-out organelles, bacteria, and other particles.

    Lysosomes contain enzymes that break down cellular debris for recycling. Section 4.7

    Put the following structures in order according to the pathway of secreted protein:

    The main function of the endomembrane system is building and modifying __________ and __________.

    Is this statement true or false? The plasma membrane is the outermost component of all cells. Explain.

    Animal cells have a plasma membrane but plant cells have a cell as their outermost component. Section 4.11

    Which of the following organelles contain no DNA?

    A nucleus protects and controls access to a eukaryotic cell's DNA. Mitochondrion have their own DNA, which is similar to bacterial DNA. Each chloroplast has two outer membranes enclosing a semifluid interior, the stroma, that contains enzymes and the chloroplast's own DNA. The Golgi body modifies peptides and lipids before sorting them into vesicles. Section 4.6, 4.7 and 4.9.

    Cytoskeletal elements called __________ form a reinforcing mesh under the nuclear envelope.

    A microfilament mesh called the cell cortex reinforces plasma membranes. Section 4.10.

    No animal cell has a ____________.

    Most prokaryotes, protists, fungi, and all plant cells secrete a wall around the plasma membrane. Many eukaryotic cells secrete a waxy, protective cuticle. Section 4.11.


    How Archaea might find their food: Sensor protein characterized

    The microorganism Methanosarcina acetivorans lives off everything it can metabolise into methane. How it finds its sources of energy, is not yet clear. Scientists at the Ruhr-Universität Bochum together with colleagues from Dresden, Frankfurt, Muelheim and the USA have identified a protein that might act as a "food sensor." They characterised the molecule in detail and found both similarities and differences to the system that is responsible for the search for food in bacteria.

    MsmS has a different function to that thought

    The protein MsmS has so far only been studied from a bioinformatics point of view. Computer analyses of its gene sequence had predicted that it might be a phytochrome, i.e. a red light sensor. Using spectroscopic methods, the research team of the current study have refuted this theory. MsmS has a heme cofactor, like haemoglobin in red blood cells, and can, among other things, bind the substance dimethyl sulphide. This is one of the energy sources of Methanosarcina acetivorans. MsmS might thus serve the microorganism as a sensor to directly or indirectly detect this energy source. In genetic studies, the scientists also found evidence that MsmS regulates systems which are important for the exploitation of dimethyl sulphide.

    Archaea: flexible "eaters"

    Methanosarcina acetivorans belongs to the Archaea which constitute the third domain of life, alongside Bacteria and Eukarya the term Eukarya comprising all living organisms with a cell nucleus. Many of them are adapted to extreme conditions or are able to use unusual energy sources. Among the organisms that live from methane production, the so-called methanogenic organisms, M. acetivorans is one of the most flexible when it comes to the choice of food sources. It converts many different molecules into methane, and thus produces energy. How M. acetivorans detects the different food sources, is still largely unknown.


    Genome-reconstruction for eukaryotes from complex natural microbial communities

    Microbial eukaryotes are integral components of natural microbial communities, and their inclusion is critical for many ecosystem studies, yet the majority of published metagenome analyses ignore eukaryotes. In order to include eukaryotes in environmental studies, we propose a method to recover eukaryotic genomes from complex metagenomic samples. A key step for genome recovery is separation of eukaryotic and prokaryotic fragments. We developed a k-mer-based strategy, EukRep, for eukaryotic sequence identification and applied it to environmental samples to show that it enables genome recovery, genome completeness evaluation, and prediction of metabolic potential. We used this approach to test the effect of addition of organic carbon on a geyser-associated microbial community and detected a substantial change of the community metabolism, with selection against almost all candidate phyla bacteria and archaea and for eukaryotes. Near complete genomes were reconstructed for three fungi placed within the Eurotiomycetes and an arthropod. While carbon fixation and sulfur oxidation were important functions in the geyser community prior to carbon addition, the organic carbon-impacted community showed enrichment for secreted proteases, secreted lipases, cellulose targeting CAZymes, and methanol oxidation. We demonstrate the broader utility of EukRep by reconstructing and evaluating relatively high-quality fungal, protist, and rotifer genomes from complex environmental samples. This approach opens the way for cultivation-independent analyses of whole microbial communities.

    © 2018 West et al. Published by Cold Spring Harbor Laboratory Press.

    Figures

    Comparison of CG_WC and CG_bulk…

    Comparison of CG_WC and CG_bulk community composition. The relative abundances of taxonomic groups…

    Identification of scaffolds for eukaryotic…

    Identification of scaffolds for eukaryotic gene prediction with EukRep. ( A ) Schematic…

    Eukaryotic gene prediction on metagenomic…

    Eukaryotic gene prediction on metagenomic scaffolds. ( A ) Gene predictions for nine…

    Overview of binned eukaryotic genomes.…

    Overview of binned eukaryotic genomes. Genomes that share greater than 99% average nucleotide…

    Phylogenetic placement of binned eukaryotic…

    Phylogenetic placement of binned eukaryotic genomes with maximum likelihood analysis of 16 concatenated…

    Comparison of CG_WC and CG_bulk…

    Comparison of CG_WC and CG_bulk metabolic capacity. Log 2 ratio of all annotated…


    Watch the video: Archaea (December 2022).