General Microbiology - Schlegel, H. 1987

The Cell and Its Structure
Prokaryotic Cell (Protocyte)

We will examine the Structure and certain Functions of Prokaryotic Cells in greater detail. Before diving into the specifics, let us highlight the primary features that distinguish a Introduction/4.html">Prokaryotic Cell from a eukaryotic one. As previously mentioned, prokaryotic cells are extremely small. The vast majority of Bacteria are rod-shaped, measuring no more than 1 µm in thickness and 5 µm in length. Many pseudomonads are 0.4–0.7 µm wide and 2–3 µm long. The diameter of micrococci is a mere 0.5 µm. True "giants" among bacteria are quite rare (such as Chromatium okenii, Thiospirillum jenense, and Achromatium), and all such forms grow relatively slowly.

Cellular compartmentalization is significantly less pronounced than in Eukaryotic cells (Fig. 2.4). The DNA is not enclosed by a nuclear membrane, and Organelles such as Mitochondria and METABOLISM/14.html">Chloroplasts are absent. The nuclear region, which appears on an ultra-thin section electron micrograph as a net-like structure composed of fine fibrils, borders directly on the ribosome-dense Cytoplasm (Fig. 2.5). In many bacteria, invaginations of The Plasma Membrane form distinct structures within the interior of the protoplast (intracellular membranes). The plasma membrane mediates energy-yielding processes such as Respiration or Photosynthesis—functions carried out by the Mitochondrial and Chloroplast membranes in eukaryotic cells.

Prokaryotic Ribosomes are smaller than Eukaryotic cytoplasmic ribosomes, belonging to the 70S type.

All the Genetic information of a protocyte is contained within a single strand of DNA—the "bacterial chromosome." In all bacteria studied to date, this DNA molecule takes the form of a closed circular thread, with a contour length ranging from 0.25 to 3 mm. Histones are absent. In addition, many bacteria have been found to harbor extrachromosomal DNA, which consists of small, likewise closed circular DNA molecules known as Plasmids. The information contained in plasmids is not essential for cell survival.

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Fig. 2.4. A. Diagram of PROKARYOTIC Cell Structure (Cytology/practical/54.html">Longitudinal section of a bacterial cell). Гли – Glycogen granules; Ж – flagellum; Кпс – capsule; КСт – Cell wall; Ли – lipid droplets; ПГМ – poly-β-hydroxybutyric acid; Пи – pili; Плз – plasmid; ПМ – plasma membrane; ПФ – polyphosphate granules; Pи – ribosomes and Polysomes; Я – nucleus (nucleoid); S – sulfur inclusions. B. Various cytoplasmic structures.

As a rule, bacteria reproduce by binary fission. The Cell elongates, followed by The formation of a transverse septum that gradually grows inward from the periphery (or via constriction), after which the daughter cells separate. However, under certain environmental conditions, the daughter cells of many bacteria remain attached to one another for some time following division, forming characteristic aggregates. Depending on the orientation of the division planes and the number of divisions, various shapes arise; for instance, spherical bacteria may form cell pairs (Diplococcus), chains (Streptococcus), sheets, or packets (Sarcina). Rod-shaped bacteria can also form pairs or chains of cells. Reproduction by budding is extremely rare among prokaryotes. Cell Division is preceded by the duplication, or Replication, of the bacterial chromosome. However, the diploid phase in the Cell Cycle is restricted to a very brief stage. Consequently, prokaryotes are haploid organisms.

Fig. 2.5. Transmission electron micrograph of an ultra-thin section of an Escherichia coli B cell. Mild plasmolysis has caused the protoplast to pull away from The cell wall, making the cell wall (КСт) and plasma membrane (ПМ) clearly distinguishable. H – nucleoid (nucleus); ЦПл – cytoplasm with ribosomes. Magnification 56,200×, inset at bottom right 216,600×. (Photo by N. Frank, Max Planck Institute for Virus Research, Tübingen.)

With few exceptions (such as Mycoplasma), prokaryotic cells are surrounded by a cell wall. The primary "framework" of the wall consists of peptidoglycan, or murein. This is a characteristic prokaryotic heteropolymer absent in eukaryotes. Many prokaryotes are motile, moving by swimming or gliding. The Organs of locomotion in swimming bacteria are specialized "bacterial" flagella. These flagella have a much simpler structure than eukaryotic ones, consisting of a single fibril.

With few exceptions, all bacteria can be classified by shape into spherical, cylindrical (rod-shaped), or curved cylindrical forms. The fundamental shapes are therefore cocci, straight rods, and Curved Rods (Fig. 2.6). Bacteria belonging to the genera Pseudomonas and Bacillus appear under the microscope as straight rods. Spirilla possess a corkscrew-like Morphology. Curved rods are referred to as vibrios (Vibrio). Certain bacteria exhibit deviations from these basic shapes. For instance, the genus Corynebacterium and coryneform bacteria typically display a club-shaped or irregularly pleomorphic morphology. Many Mycobacterium species show traces of branching. Streptomycetes even form a mycelium resembling that of Fungi, though distinguished from the latter by a smaller cell diameter (less than 1 µm, compared to over 5 µm in fungi).

Fig. 2.6. Morphologies of unicellular bacteria. 1 – micrococci; 2 – diplococci; 3 – streptococci; 4 – staphylococci; 5 – sarcinae; 6 – rod-shaped bacteria; 7 – spirilla; 8 – vibrios.

Chemical composition of unicellular organisms. The wet biomass of bacteria is determined after separating the cells from the liquid growth medium via centrifugation. The pelleted cell mass contains 70–85% Water, meaning dry biomass accounts for 15–30% of the wet weight. If the cells contain large amounts of reserve Materials (Lipids, Polysaccharides, polyphosphates, or sulfur), the proportion of dry mass is higher. Bacterial dry matter consists predominantly of polymers [Proteins (50%), cell wall components (10–20%), RNA (10–20%), DNA (3–4%)], alongside lipids (10%). The ten most abundant chemical elements are represented in bacterial cells approximately as follows: carbon – 50%, oxygen – 20%, nitrogen – 14%, hydrogen – 8%, phosphorus – 3%, sulfur – 1%, potassium – 1%, calcium – 0.5%, magnesium – 0.5%, and iron – 0.2%.

2.2.1 The Bacterial Nucleus

The small size of the bacterial cell and the presence of two Types of Nucleic acids made the cytochemical detection of nuclear material extremely difficult. Nevertheless, classical cytological Methods, followed by ultra-thin sectioning techniques combined with Electron Microscopy, ultimately established that bacteria contain DNA, and that this DNA is not diffusely distributed throughout the cytoplasm but rather localized in discrete regions that divide prior to cell division.

Cytological detection of the bacterial nucleus. The ability of DNA to undergo specific staining underlies the Feulgen reaction for nuclear matter. The interaction between free aldehyde groups and colorless sulfurous acid-fuchsin produces a purple color characteristic of basic fuchsin. To remove RNA from the cells and liberate the aldehyde groups of the deoxypentose component of DNA, the cells are pretreated with hot dilute HCl (4 min in 1 M HCl at 60°C). Optimal results were obtained by staining the cells with the basic dye Giemsa following identical pretreatment. Fixation methods were subsequently refined; RNA began to be removed using Ribonuclease. This approach yielded preparations of Escherichia coli, Bacillus cereus var. mycoides, and B. megaterium that led to the Conclusion that bacteria possess distinct structures resembling a nucleus, referred to as Chromatin bodies, nucleoids, or nuclear equivalents (Figs. 2.7 and 2.8).

Fig. 2.7. Bacterial nuclear structures. A. Bacillus cereus. B and C. Micrococcus radiodurans, strain "Sark". The first two images are bright-field micrographs following Giemsa staining; the third image utilizes phase contrast. (Photos by C. F. Robinow.)

Progress in determining The structure of the bacterial nucleus was achieved only through transmission Electron Microscopy of ultra-thin sections of bacterial cells. Proper fixation (using osmium tetroxide, uranyl acetate, or phosphotungstic acid) under strictly controlled conditions was crucial for obtaining an optimal depiction of the native Fine Structure of the Cell Nucleus. The nuclear region (nucleoplasm) in a bacterial cell is uniformly filled with exceptionally fine fibrils (Fig. 2.5). In the Electron microscope, it appears less dense than the surrounding ribosome-rich cytoplasm. No Membrane Structure separating the nuclear region from the cytoplasm could be detected.

Autoradiography. Cairns was the first to demonstrate—using autoradiography—that the nuclear material consists of DNA and, in Escherichia coli for example, is represented by a single closed circular strand approximately 1 mm long. To achieve this, preparations of cells grown in a medium containing tritium-labeled thymidine (a precursor of thymine) (³H) were used. DNA is the sole substance in the cell containing thymine. When cells that have incorporated ³H into thymine are lysed with Lysozyme or sodium lauryl sulfate on a membrane filter, an autoradiograph of the uncoiled bacterial chromosome can be obtained. Such autoradiographs (Fig. 2.9) provide compelling evidence that bacterial DNA takes the form of a closed circular thread. Genetically, this thread corresponds to a linkage group and is designated as the bacterial chromosome. Furthermore, autoradiographs offer insight into the replication mechanism of such a thread. Fig. 2.10 depicts bacteriophage DNA visualized via an alternative method.

Fig. 2.8. ELECTRON MICROGRAPHS OF enzyme-treated Escherichia coli cells. In the cells shown in the top micrograph, the cytoplasm and its RNA have been digested by ribonuclease and Pepsin, leaving only the cell walls and nucleoids (nuclei) intact. The cells in the bottom micrograph were treated sequentially with ribonuclease, deoxyribonuclease, and pepsin, resulting in the degradation of the nucleoids as well, leaving only empty cell walls. This sequential differential enzymatic Treatment demonstrates that the dark regions (in the top micrograph) correspond to nucleoids (DNA). (Peters D., Wigand R., Z. Naturforsch., 8b [1953], 180.)

Fig. 2.9. Autoradiographs of replicating Escherichia coli Chromosomes. The cells were grown for two generations in the presence of tritium-labeled thymine and subsequently lysed with lysozyme; for details, see text. (Bleeckens S., Strohbach G., Sarfert E., Z. allg. Microbiol., 6 [1966], 121.)

DNA Structure. Deoxyribonucleic acid (DNA) is a macromolecule. Upon acid Hydrolysis, it breaks down into its fundamental structural units—deoxyribose, phosphoric acid, and nitrogenous bases in equimolar ratios. DNA contains four different bases: two Purines (adenine and guanine) and two Pyrimidines (cytosine and thymine).

Fig. 2.10. PM2 bacteriophage DNA molecules shown in various states. Each tangle consists of ccc-DNA (covalently closed circular DNA). Ultraviolet irradiation caused single-strand breaks in some molecules, producing oc-DNA (open circular DNA) that appears as a large ring with a contour length of 3.02 µm. The DNA was spread using a quaternary ammonium compound and stained with uranyl acetate (positive contrast); dark-field electron micrograph. (U. Hahn, Göttingen.)

Digesting DNA with Nucleases (such as pancreatic DNase I or snake venom diesterase) releases 3'- or 5'-NUCLEOTIDES. In Nucleic Acids, these deoxyribonucleotides are linked into long chains. In DNA, pentose molecules alternate with phosphoric acid molecules, with one of the four bases attached to each sugar. Such a polynucleotide chain is asymmetric—that is, it exhibits polarity: one end features a 5'-phosphate group, while the other bears a free 3'-hydroxyl group.

As early as 1950, Chargaff established several key regularities: The amount of adenine in DNA equals that of thymine, and the amount of guanine equals that of cytosine (A = T, G = C); the sum of purine bases equals the sum of pyrimidine bases; and the (G + C)/(A + T) ratio varies widely among different species but remains constant for any given species.

The spatial arrangement of the individual components (i.e., the Secondary structure of DNA) was elucidated using X-Ray Diffraction Analysis. When a DNA fiber is stretched and exposed to a monochromatic X-ray beam, projecting the deflected rays onto an emulsion film yields an X-ray diffraction pattern. Diffraction patterns of DNA from various sources (sperm, Thymus, bacterial cells, and Bacteriophages) proved nearly identical. Wilkins's analysis of these patterns demonstrated that the purine and pyrimidine rings lie at right angles to the longitudinal axis of the polynucleotide strand, which must be helically coiled around a central axis with a pitch of 3.4 nm. Density data further suggested that the helix comprises more than a single strand.

These findings enabled Watson and Crick to formulate their brilliant theory of DNA structure in 1953. According to the Watson-Crick model, the polynucleotide chains are wound into a double helix around an imaginary axis. The two chains are held together by Hydrogen Bonds between the bases, which face inward toward the center of the helix (Fig. 2.11). Accounting for interatomic distances and bonding constraints, adenine must always pair with thymine, and guanine with cytosine. Each turn of the helix contains approximately 10 Base Pairs. The base sequences of the two strands are consequently complementary, and the strands run in opposite directions (5' → 3' and 3' → 5'). The chromosome of Escherichia coli is roughly 1.4 mm long. One micrometer of single-stranded DNA corresponds to a relative molecular mass of about 2·10⁶, or approximately 3,000 base pairs. Thus, the E. coli chromosome has a molecular mass of 2.9·10⁹.

The hydrogen bonds connecting A to T and G to C vary in strength and are predominantly electrostatic in nature. They involve OH and NH₂ groups. Because oxygen and nitrogen are strongly electronegative elements, they attract electrons and impart a partial positive charge to the hydrogen atoms bound to them. A positively charged hydrogen atom can then be attracted by other electronegative groups possessing lone electron pairs, forming a Hydrogen bond. Its strength depends on the acidity of the hydrogen atom and the basicity of the acceptor atom. Hydrogen bonds are stronger than Van der Waals forces, with a bond energy reaching up to 38 kJ (9 kcal) per mole—on average only slightly greater than the thermal agitation energy at 37°C. As shown in Fig. 2.11, three hydrogen bonds link guanine and cytosine, while two connect thymine and adenine. Because the bond energy is low, factors such as elevated temperatures, minor shifts in magnesium concentration, or The addition of urea can cause significant conformational changes or even strand Separation. Raising the Temperature breaks the Hydrogen bonds and causes the polynucleotide strands to dissociate. This disruption of the DNA secondary structure is accompanied by an increase in Light absorption at 259 nm (the hyperchromic effect). The temperature at which the absorbance increase reaches half its maximum value is termed the melting point (Tm). The melting point increases with the proportion of guanine and cytosine, which are joined by three hydrogen bonds. Consequently, the melting point of isolated, purified DNA (Fig. 2.12) serves as a convenient indicator for determining its relative GC content. The GC content is expressed as the percentage of the sum of guanine and cytosine moles relative to the total moles of all four bases in the given DNA.

Fig. 2.11. Structure of deoxyribonucleic acid (DNA). The single DNA strands are joined by hydrogen bonds. Left: the DNA double helix, shown as a space-filling model at the top, and with the base pairs arranged like the steps of a spiral staircase below. Right: base pairing of adenine with thymine and guanine with cytosine. Dashed lines represent hydrogen bonds.

Fig. 2.12. A. Model of a plectonemic double helix, which can be formed by twisting two parallel wires around a rod and then withdrawing the rod. The two single strands are intertwined and can only be separated by uncoiling. B. Melting curve of bacterial DNA. Tm is the temperature at which absorbance reaches half its maximum value.

Bacterial DNAs vary widely in their GC content, which ranges from 30% (in certain staphylococci and members of the Cytophaga group) to over 70% (in Representatives of the genus Micrococcus and certain fruiting myxobacteria). GC content is species-specific and serves as an important taxonomic marker.

DNA replication. DNA carries the genetic information of the cell. Therefore, the duplication of DNA—which invariably precedes cell division—must yield two identical chromosomes. At first glance, this process (identical reduplication or replication of DNA) appears very straightforward: the two strands simply need to separate so that free nucleotides can align opposite their complementary bases along each single polynucleotide strand, followed by polymerization into a new chain. One of the primary conceptual challenges is understanding how the two parental strands unwind. X-ray diffraction data reveal that DNA forms a plectonemic rather than a paranemic double helix (Fig. 2.12, A). To determine whether unwinding is actually required, three theoretically possible mechanisms of DNA replication were evaluated as working hypotheses (Fig. 2.13) (Delbrück and Stent, 1957):

1. Conservative mechanism. The Double Helix does not unwind; the parental double helix acts as a template for synthesizing two entirely new strands. The daughter double helix is built completely from fresh material, while the parental helix remains intact.

2. Dispersive mechanism. During duplication, the parental helix undergoes extensive fragmentation every half-turn; new strands are synthesized on these fragments, which then recombine crosswise with segments of new material. Each resulting polynucleotide chain would thus consist of alternating segments of old and new material.

3. Semiconservative mechanism. The parental double helix unwinds, and a new complementary strand is synthesized along each separated polynucleotide chain. Consequently, each new double helix forms a "hybrid" consisting of one old and one newly synthesized strand.

Fig. 2.13. Left: model of semiconservative DNA replication. Right: schematic representations of the three theoretically possible mechanisms of DNA replication: conservative (1), dispersive (2), and semiconservative (3). Explanations are provided in the text.

To ascertain which of these three hypotheses reflects reality, Meselson and Stahl performed labeling experiments coupled with density gradient centrifugation of the labeled DNA (Fig. 2.14).

Centrifuging a 6 M cesium chloride solution for many hours at 100,000 g establishes a density gradient (at equilibrium between centrifugal force and diffusion). If the solution contains DNA, it concentrates in a zone whose position in the gradient corresponds to its buoyant density. DNA extracted from bacteria grown in the presence of ¹⁵NH₄Cl is about 0.8% heavier than normal ¹⁴N-DNA, causing it to form a distinct band in the CsCl gradient. Escherichia coli bacteria were cultured for numerous generations with ¹⁵N as the sole nitrogen source, ensuring their DNA contained exclusively ¹⁵N. Subsequently, an excess of ¹⁴N was added to the growth medium. Samples were taken before and after this addition, and the extracted DNA was analyzed in a CsCl density gradient. After one cell division in the ¹⁴N medium, all the DNA exhibited an intermediate density ("semi-heavy" or "hybrid" DNA). The total amount of this semi-heavy DNA remained constant over several generations, whereas the amount of light DNA increased. To prove that one strand of the semi-heavy DNA indeed contained ¹⁵N and the other ¹⁴N, the semi-heavy DNA was "melted" (heated to 100°C) and rapidly cooled. Subsequent CsCl density gradient centrifugation revealed two bands: one characteristic of single-stranded ¹⁴N-DNA, and the other of single-stranded ¹⁵N-DNA.

Fig. 2.14. Semiconservative DNA reduplication. Outline of the Meselson-Stahl experiment. Separation of ¹⁵N- and ¹⁴N-labeled DNA by centrifugation in a 6 M CsCl solution. 1 - density gradient formed by equilibrium between diffusion and sedimentation forces; 2 - DNA from normal cells grown in ¹⁴NH₄Cl medium; 3 - DNA from cells grown for several generations in ¹⁵NH₄Cl medium; 4 - DNA from ¹⁵N-labeled cells grown for one generation in ¹⁴NH₄Cl medium; 5 - DNA from ¹⁵N-labeled cells grown for two generations in ¹⁴NH₄Cl medium; 6 - DNA from ¹⁵N-labeled cells grown for three generations in ¹⁴NH₄Cl medium.

The results of these experiments are incompatible with either the conservative or dispersive mechanisms of DNA replication; instead, they fully support the semiconservative model. During replication, the strands unwind and separate, and a new complementary strand is synthesized along each single template strand. DNA polymerases mediate this process, functioning simply to link together nucleotides that have aligned via base pairing in the correct sequence, thereby building a new polynucleotide chain. The biochemical mechanisms are illustrated in Fig. 2.15 and detailed in its caption. Further details can be found in textbooks on molecular biology.

Synthesizing one polynucleotide chain in this manner is straightforward: DNA polymerase can continuously link nucleotides in the 5' to 3' direction. However, because the DNA double helix is antiparallel, Synthesis of the complementary, opposite strand must proceed in the reverse direction. These theoretical and experimental insights led to the model illustrated in Fig. 2.16. Initially, short segments approximately 1,000 nucleotides long—known as Okazaki fragments—are synthesized. Each fragment begins with the formation of a short RNA oligonucleotide that serves as a primer. A DNA polymerase then extends this RNA primer with a DNA segment 1,000 to 2,000 nucleotides long. Finally, the RNA primer is excised, the resulting gap is filled by another polymerase, and the adjacent segments are joined together by DNA ligase. This discontinuous synthesis mechanism successfully explains the replication of the second DNA strand, and it is even possible that both strands replicate in this fashion.

Fig. 2.15. Functioning of DNA polymerases. The DNA double helix consists of two polynucleotide chains of opposite polarity (they are "antiparallel"). If the "free" 3'-OH group, not bound to an adjacent nucleotide, is located at the left end of one chain, an identical group is found at the right end of the other chain. DNA replication is catalyzed by DNA polymerases. The functioning of such Enzymes requires: 1) a template, which is a single strand of DNA; 2) a primer, a short segment of replicated nucleic acid; and 3) a mixture of deoxynucleoside 5'-triphosphates. DNA polymerases can attach free nucleotides exclusively to the free 3'-OH end of a nucleotide chain. Thus, synthesis proceeds exclusively in the 5' → 3' direction and not vice versa.

Fig. 2.16. Replication of double-stranded DNA. During DNA uncoiling, an intermediate branched structure (the Replication fork) is formed. The two single DNA strands possess opposite polarity (5' → 3' and 3' → 5'). DNA polymerases are capable of catalyzing synthesis in only one direction (5' → 3'). Therefore, the synthesis of one strand can proceed continuously in the direction of the advancing unwinding of the double helix. The other strand, however, must be synthesized in the reverse direction. Synthesis begins with the formation of a short RNA segment acting as a "primer" (A-C-S-A bases in bacteriophage T7). Subsequently, DNA polymerase synthesizes a DNA strand 1,000–2,000 nucleotides long adjacent to this RNA. Eventually, the RNA primer is removed by an exonuclease, the gap is filled by DNA polymerase (a), and sealed by DNA ligase (b). Such a mechanism of "fractional" or "discontinuous" DNA Synthesis, followed by the joining of individual segments, explains DNA replication on the antiparallel strand.

Fig. 2.17. Replication of a circular bacterial chromosome and Bacterial cell division. This diagram assumes a unidirectional mechanism of DNA replication, i.e., involving only a single replication fork.

Fig. 2.17 illustrates how one can conceptualize the replication of a bacterial chromosome and bacterial cell division; this diagram assumes that the replication of the entire DNA ring proceeds in a single direction.

However, it has been found that the synthesis of both new strands can proceed not only in a single direction but also bidirectionally from the initiation point. Such a mechanism involves the unwinding of the double helix at two sites simultaneously, resulting in the formation of two branching replication forks on a single DNA molecule. Chromosome duplication in E. coli takes approximately 40 minutes. Meanwhile, under favorable conditions, this bacterium divides with a doubling time of only about 20 minutes. This fact can be explained by data indicating that both daughter chromosomes initiate a new round of division even before the previous one is completed. Much more detailed information on DNA Replication Mechanisms has been obtained than can be presented here. There are significant differences among the replication mechanisms of phages, plasmids, and bacterial chromosomes. For an in-depth study of these topics, one should consult literature on molecular biology.

Genome sizes and number of genomes. The Genome Size in bacteria varies from species to species, ranging from 0.8 to 8×10⁶ base pairs. The number of genomes per cell also varies among species and further depends on cultivation conditions. In batch culture, growing cells of Escherichia coli contain 2 to 4, Azotobacter chroococcum 20 to 25, and Desulfovibrio gigas 10 to 15 genomes per cell. (For comparison, eukaryotic genome sizes are: Neurospora crassa 19×10⁶, Aspergillus niger 40×10⁶, human 2.9×10⁹, and Zea mays 7×10⁹ base pairs.)

DNA/DNA Hybridization: homologous nucleotide sequences in DNA of different species. As noted earlier, when isolated DNA is heated, the two polynucleotide chains separate due to the rupture of hydrogen bonds. This Denaturation (or "melting"), leading to the formation of single strands, is reversible: upon very slow cooling of the preparation, pairing and reassociation of complementary regions will take place. If short fragments of denatured DNA obtained from two different yet closely related bacterial species are mixed at a temperature above their melting point and the mixture is then slowly cooled, reassociation will also occur. Double helices formed from single DNA strands of two different organisms are called heteroduplex molecules. To experimentally track the formation of heteroduplexes, the DNA of one of the bacteria must, of course, be labeled with a heavy or radioactive isotope.

If, for example, a culture of one bacterium is grown in heavy water (D₂O), it will yield "heavy" DNA, and the Formation of the heteroduplex can then be detected by CsCl density gradient centrifugation—just as the formation of ¹⁴N/¹⁵N hybrid DNA was proven in the Meselson-Stahl experiment. Alternatively, the DNA of one partner can be labeled by growing it on a medium containing ¹⁴C or ³²P. If long stretches of denatured DNA (from unlabeled ¹²C [sic] bacterium A) are mixed with short stretches of denatured DNA (from ¹⁴C-labeled bacterium B), the mixture is slowly cooled and passed through a filter that retains long chains while letting short ones through, heteroduplex molecules will remain on the filter. The radioactivity remaining on the filter will be higher the more radioactive segments (B) have bound to the A-strands, i.e., it will depend on whether the base sequences of DNA A and B are identical (or very similar) or completely different. Thus, DNA/DNA reassociation makes it possible to determine the degree of Sequence Homology of DNA from different origins. A control experiment is performed using DNA molecules from the same bacteria (one sample labeled, the other unlabeled), and the degree of reassociation is arbitrarily set at 100. The degree of reassociation of DNA molecules from various strains is then expressed as a percentage of this value. Routine Methods for determining homology between DNA of various bacterial strains have been modified many times, but all are based on the same principle of heteroduplex formation; in all cases, labeling of one of the partners is required.

DNA homology, i.e., the degree of base sequence identity in DNA molecules of two different bacterial strains, is greater the closer the phylogenetic relationship between these strains. This method has proven reliable when comparing closely related strains and species; however, between distantly related genera, the degree of DNA homology is too low to detect heteroduplex formation. Single-stranded DNA can also reassociate with RNA. Therefore, the described methods also allow the assessment of homology between DNA and RNA.

Plasmids. Alongside chromosomal DNA, many bacteria contain extrachromosomal DNA, which also consists of double-stranded closed rings. These autonomously replicating DNA elements are called plasmids (see the subsection "Plasmids" in Section 15.3.2).

2.2.2 Cytoplasm and membranes

The cytoplasm is separated from the cell wall by the plasma membrane. The cytoplasm contains various inclusions (vesicles, granules) and the nucleoid. As electron-microscopic and biochemical studies have shown, the cytoplasm is not a homogeneous protein solution; it contains numerous membranes and various membrane structures, while the remaining space is occupied by the liquid phase and ribosomes. Prolonged centrifugation at 100,000 × g can separate an aqueous suspension of cytoplasm into a "soluble" fraction, containing primarily soluble enzymes and soluble ribonucleic acid (RNA), and a particulate fraction, which, along with membranes, predominantly comprises ribosomes. Soluble enzymes catalyze numerous Catabolic and anabolic reactions. Soluble Ribonucleic Acids [messenger (mRNA) and transfer (tRNA)] and ribosomes participate in Protein Synthesis.

Proteins. Proteins consist of Amino Acids linked in a specific sequence by peptide bonds into polypeptide chains. These chains possess a strictly defined spatial configuration (conformation) stabilized by additional bonds—covalent and non-covalent (Fig. 2.18). Depending on the role played by particular bonds in maintaining structure, several structural levels are distinguished. The Primary Structure of a protein is determined by the number and sequence of covalently linked amino acids. Through hydrogen bonds formed between oxygen atoms of carbonyl groups and nitrogen atoms of amide groups, the polypeptide chain acquires secondary structure; it can form a helical configuration (α-Helix) or a so-called pleated sheet configuration. Tertiary structure refers to the specific spatial arrangement of the peptide chain resulting from interactions among its various side groups. Other hydrogen bonds, ionic bonds, and non-polar (hydrophobic) interactions participate in maintaining the tertiary structure. Cross-links connecting different Regions of the polypeptide chain can also be covalent; Examples include Disulfide Bonds formed by The oxidation of SH groups. Finally, supermolecular aggregates can arise through interactions among multiple polypeptide chains. This architecture (in which a protein consists of a specific number of polypeptide chains, or subunits) is called quaternary structure. Under physiological conditions, proteins exist in an aqueous phase. Therefore, interactions also occur between proteins and water dipoles. Polar groups are hydrated. Factors that alter the charge of proteins (concentrations of H⁺, Ca²⁺, Mg²⁺, K⁺ ions, etc.) inevitably affect the degree of Hydration and, consequently, the degree of protein Swelling.

Fig. 2.18. Possible intramolecular bonds between different regions of a polypeptide chain. a — covalent disulfide bonds; b — hydrogen bonds; c — non-polar (hydrophobic) interactions; d — ionic bonds. [Lynen F., Naturw. Rdsch., 23, [1970], 266.]

Ribosomes. Ribosomes serve as the site of protein synthesis. In electron micrographs, they appear as particles situated within the cytoplasm. Bacterial ribosomes measure 16×18 nm. Approximately 80–85% of total bacterial RNA is located in ribosomes. Because intact bacterial ribosomes sediment during ultracentrifugation at a rate of about 70 Svedberg units (S), they are called 70S ribosomes. Eukaryotic cytoplasmic ribosomes, with few exceptions, are somewhat larger and are designated as 80S ribosomes.

Ribosomes consist of two subunits; in bacteria, these are the 30S and 50S subunits (Fig. 2.19), which assemble into 70S ribosomes. In size and many other features, bacterial ribosomes resemble mitochondrial and chloroplast ribosomes. A bacterial cell contains roughly 5,000 to 50,000 ribosomes; their number increases with the growth rate of the cell. During active protein synthesis, regular chains of ribosomes can be observed in electron micrographs of thin cell sections. These are ribosomes strung like beads along an mRNA chain, known as polyribosomes or polysomes.

Differences between bacterial (70S) and eukaryotic (80S) ribosomes are of decisive importance for combating infectious diseases; certain Antibiotics partially or completely inhibit protein synthesis occurring on 70S ribosomes without affecting the function of 80S ribosomes (Section 6.6).

Fig. 2.19. Bacterial ribosomes. Left: electron micrograph of ribosomes from E. coli. Intact 70S ribosomes are marked with three triangles, 50S subunits with two, and 30S subunits with one. Scale bar, 100 nm. (Photo by E. Spiess.) Right: diagram of ribosome assembly from subunits and their compositional data.

RNA Structure. Ribonucleic acids differ from DNA both in their constituent monomers and in their secondary structure. The backbone of the polynucleotide chain consists of ribose and phosphoric acid. The same bases are present—adenine, guanine, and cytosine—except that uracil replaces thymine. Additionally, RNA contains several rare bases (e.g., pseudouracil). RNA is present in cells in a single-stranded form; base pairing occurs only in restricted regions of the chain.

Plasma membrane. In electron micrographs of ultrathin sections of bacteria fixed with osmium tetroxide, the plasma membrane appears multilayered. It consists of two osmiophilic and thus dark layers, each 2–3 nm thick, and an intermediate lighter layer 4–5 nm thick. In their structural Organization, the membranes of bacterial, animal, and plant cells are very similar. This provides a basis for THE CONCEPT OF the universal "unit membrane".

Membranes can be isolated by subjecting protoplasts, obtained using lysozyme, to osmotic Shock. The membrane is rich in lipids, especially Phospholipids (Table 2.1). Accounting for only 8-15% of the dry weight of the cell, membranes contain 70-90% of all its lipids.

The plasma membrane consists of a lipid bilayer. The hydrophobic tails of the phospholipid and triglyceride molecules are oriented inward, while the hydrophilic "heads" face outward. Due to hydrophobic interactions between the fatty acid residues that make up the lipids, and Electrostatic Interactions between the hydro-

Table 2.1. Composition of membranes in Micrococcus luteus (lysodeikticus) and phototrophic bacteria

philic "heads", the membrane is stabilized. Proteins are embedded within The Lipid Bilayer—these are the so-called integral Membrane Proteins. They "float" in this layer, being partially immersed in it or penetrating it entirely. Other proteins are attached to The surface of the membrane and are called peripheral proteins (Fig. 2.20). Some membranes appear to be covered on one or both sides by a network of elongated protein molecules.

The membrane should be viewed as a very soft, plastic, almost liquid structure; isolated membranes tend to form completely closed vesicles, and membrane fragments fuse at their edges with one another.

The plasma membrane plays a crucial role in metabolism. It serves as the osmotic barrier of the cell and controls both the uptake of substances into the cell and their efflux. The membrane features Active Transport MECHANISMS and substrate-specific permease systems. It appears that the lipid film of the unit membrane is pierced by "bridges" (or channels) made of proteins, and it is these proteins that serve as pores through which regulated Transport of substances takes place.

The enzymes of electron transfer and Oxidative Phosphorylation, which in eukaryotes reside in the mitochondria, are localized in bacteria within or On the surface of the plasma membrane. Cytochromes, iron-sulfur proteins, and Other components of the Electron Transport Chain are found exclusively in the membranes. As detailed studies of the localization of individual components have shown, the membrane has an asymmetrical structure: for example, cytochrome c is located in its outer layer, while ATP synthase is on the inner side of the membrane. The membrane is characterized by directed metabolism (Sec. 7.4). Membranes are presumably also responsible for other biosynthetic processes, such as the Synthesis of cell wall and capsule components, as well as The excretion of extracellular enzymes. Finally, it is very likely that the RNA replication center is localized on the membrane. Flagella are also attached to the plasma membrane.

Fig. 2.20. Model of the plasma membrane structure. Integral proteins are embedded in the lipid bilayer. Peripheral proteins merely adhere to the membrane surface.

Fig. 2.21. Mesosomes. A. Lactobacillus fermentum, electron micrograph of an ultrathin section after fixation with osmium tetroxide and post-staining with uranyl acetate. (Photo by F. Schötz.) B. Lactobacillus viridescens (syn. corynoides) with densely packed tubules lying parallel or perpendicular to the plane of the section, (F. Schötz, J. G. Abo-Elnaga, O. Kandler, Z. Naturforsch., 20 [1965], 790.) C. Unidentified bacterium (strain 11x), electron micrograph, negative contrast (phosphotungstic acid). (Photo by R. Hillmer, F. Amelunxen.)

Intracellular membranes and lamellae. In some bacteria, the membrane encloses the cytoplasm without folds or invaginations. In others, it forms invaginations, penetrates the cytoplasm, or gives rise to membrane bodies. Mesosomes have been described in A number of bacteria (Fig. 2.21). However, in this case, these are likely artifacts associated with specimen preparation.

Nitrobacter, Nitrosomonas, and Nitrosococcus possess stacks of lamellae consisting of parallel flattened vesicles, some of which are connected to the plasma membrane (Figs. 2.22 and 2.24).

Phototrophic purple bacteria are particularly rich in intracellular membranes. In ultrathin sections, their internal photosynthetic membrane systems appear as tubes, vesicles, or stacks. In Rhodospirillum rubrum or Chromatium, the cell is almost entirely filled with vesicles packed into rounded structures (Fig. 2.23). These vesicles apparently arise through the invagination of the plasma membrane and its growth in the form of sleeves. Constrictions form at regular intervals along this sleeve, resulting in vesicles that are not completely separated from one another. Only upon Cell Disruption and homogenization are these vesicles released, taking the form of isolated structures formerly sometimes referred to as "chromatophores." In other purple bacteria, the vesicles are strongly flattened and form ordered stacks (Figs. 2.23, 2.24); by analogy with the Structure of Chloroplasts in green plants, the latter are called thylakoid stacks.

Fig. 2.22. Electron micrographs of the nitrifying marine bacterium Nitrosococcus oceanus: A and C - ultrathin sections (after OsO₄ fixation, 22,000x), B - freeze-etch preparation (81,000x). (Remsen C. C., Valois F. W., Watson S. W., J. Bacteriol. 94 [1967], 422.) CW - cell wall; PM - plasma membrane; DL - double lamella.

Fig. 2.23. Photosynthetic Pigments in phototrophic purple bacteria are associated with intracellular membranes. The expansion of membrane surface area in various bacterial species is achieved in different ways. In Chromatium okenii, these structures appear as vesicles and are called vesicles or chromatophores (A). Thiocapsa pfennigii contains tubular photosynthetic membranes (B). In Ectothiorhodospira mobilis, the membranes are folded multiple times to form lamellar stacks (C). CW - cell wall; PG - polysaccharide granules; LS - lamellar stacks; TM - tubular photosynthetic membranes;

Fig. 2.24. Ectothiorhodospira mobilis with stacks of photosynthetic lamellae. Electron micrograph of an ultrathin section (150,000x). PM - plasma membrane; R - ribosomes; LS - lamellar stacks; 1 - outer layer of the cell wall; 2 - inner layer of the cell wall; 3 and 4 - outer and inner electron-lucent intermediate layers.

Photosynthetic membranes are similar in Structure and Chemical composition to the plasma membrane (Table 2.1), but in addition contain light-absorbing pigments (bacteriochlorophylls and carotenoids) as well as Components of the photosynthetic electron transport chain (cytochromes, ubiquinone) and the phosphorylating system.

2.2.3 Cell Wall

The Bacterial cell wall is not rigid like a steel armor, but thin and elastic like the leather cover of a football. Just as a ball is kept firm by an inflated inner bladder, the cell wall is given a certain elasticity by the protoplast tightly pressed against it from within. Internal pressure (turgor) is driven by osmotic factors. The plasma membrane serves as the osmotic barrier: it is semipermeable and controls the entry and exit of dissolved substances. Unlike the plasma membrane, the cell wall is permeable to salts and other low-molecular-weight compounds.

Plasmolysis. Under normal conditions, the concentration of sugars and salts—osmotically active substances—inside the cell is higher than in the external environment. In terms of osmotic pressure, the cell contents are equivalent to a 10-20% sucrose solution, and water enters the cell to the extent allowed by the stretchability of its wall. If the Osmotic Pressure of the external medium is increased (e.g., by adding sugars or urea), water will be drawn out of the cell. Eventually, the protoplast will shrink and the plasma membrane will detach from the cell wall. This process, occurring in a hypertonic environment, is called plasmolysis. It is precisely The phenomenon of plasmolysis in large bacterial cells that allows us to verify that their plasma membrane is surrounded by a cell wall. Both the plasma membrane and the cell wall are stained by the water-soluble basic dye Victoria Blue.

Gram staining. The cell wall appears to be largely responsible for the Gram-staining reaction as well. The ability or inability to retain a dark purple color using the technique introduced by Gram in 1884 serves as a crucial taxonomic criterion that correlates with other bacterial properties. The Gram-Staining Procedure begins by treating fixed bacterial cells with the primary stain, crystal violet, followed by an iodine solution. Iodine forms a water-insoluble complex with crystal violet that dissolves only poorly in alcohol and acetone. The cells are then "differentiated" by washing with alcohol: Gram-positive cells retain the dye-iodine complex and remain blue, whereas Gram-negative cells are decolorized. To make them visible, the latter are counterstained with safranin or fuchsin.

If Gram-positive cells are treated with lysozyme after Gram staining, the protoplasts remain stained, but are decolorized upon subsequent exposure to alcohol. Germinating Bacillus subtilis spores and the first cell generations immediately following spore germination behave as Gram-negative organisms, only later becoming Gram-positive. These observations indicate that the staining complex resides within or on the surface of the protoplast, and that in Gram-positive bacteria it is primarily the cell wall that retains the complex by preventing its extraction.

The basic "framework" of the bacterial cell wall. To understand the STRUCTURE OF THE bacterial cell wall, it is first important to note the similarity between its supporting "Skeleton" and the polymers of ß-D-glucose, namely Cellulose and Chitin.

Cellulose is the primary component of cell walls in higher plants, Algae, and oomycetes. Cellulose does not occur as a cell wall material in bacteria, but it binds Sarcina ventriculi cells into large packets. In addition, Acetobacter aceti subsp. xylinum secretes cellulose into the medium in the form of fine fibrils, giving the "vinegar fungus" pellicle ("Mycoderma aceti") a tough, leather-like consistency.

Chitin is the structural material of the exoskeleton in Arthropods and certain other animals. It is also a vital cell wall component in Major fungal groups (Basidiomycetes, ascomycetes, and zygomycetes). The structural units of chitin are N-acetylglucosamine residues linked together analogously to the glucose residues in cellulose, i.e., via ß-1,4-glycosidic bonds.

The supporting Skeleton of the bacterial wall also consists largely of a homogeneous polymer: the peptidoglycan murein. This macromolecule is a heteropolymer built of chains in which alternating residues of N-acetylglucosamine and N-acetylmuramic acid (N-acetylglucosamine lactate) are joined by ß-1,4-glycosidic bonds.

These unbranched heteropolymeric chains form the backbone of murein. The muramic acid residues are linked via their lactyl groups to amino acids through peptide bonds. Typical amino acids of murein include L-Alanine, D-glutamic acid, meso-diaminopimelic acid or L-Lysine, and D-alanine. The diamino acids meso- (or LL-) diaminopimelic acid and L-lysine play a major role in intermolecular cross-linking, as they form peptide bonds utilizing both amino groups and can thus link two heteropolymeric chains together (Fig. 2.25). Diaminopimelic acid or lysine can sometimes be replaced by Ornithine or diaminobutyric acid. Through these peptide bridges, the heteropolymeric chains are interconnected into a sac-like giant molecule known as the murein sacculus.

Fig. 2.25. Structure of the murein sacculus in Escherichia coli. The heteropolymeric chains, consisting of alternating residues of N-acetylglucosamine (N-GlcNAc) and N-acetylmuramic acid (N-MurNAc), are interconnected by peptide bridges. Left: detailed structure of the fragment boxed on the right. Triangles indicate bonds cleaved by lysozyme (muramidase) and specific muroendopeptidases. Bottom right: schematic perspective view of the structure of a single-layer, cross-linked murein sacculus composed of N-GlcNAc (G) and N-MurNAc (M). m-Dpm — meso-diaminopimelic acid.

The classical model proposing that these chains form closed rings arranged like hoops around rod-shaped bacteria or perpendicular to them is clearly untenable. Rather, the length of the heteropolymeric chains is likely no greater than one-tenth of the bacterial circumference. Apparently, chains consisting of 50 to 500 disaccharide units (N-acetylglucosamine + N-acetylmuramic acid) are cross-linked by peptide bridges to form a less regular network than previously assumed.

Of particular note is the fact that the bacterial wall contains structures and substances absent in animals and plants, such as the alternating sequence of N-acetylglucosamine and N-acetylmuramic acid; meso-diaminopimelic acid, which does not occur in proteins; and the D-isomers of alanine and glutamic acid. These structural elements constitute the Achilles' heel of bacteria, exploited by physicians in combating infections. In terms of composition, cell wall structure, and the Biochemical Mechanisms of its synthesis, bacteria differ fundamentally from animals and plants. Therefore, drugs that specifically target only bacterial walls and their synthesis pathways remain harmless to higher organisms.

The presence of a peptidoglycan layer in cell walls is a characteristic feature of all prokaryotes, with the sole exception of archaea and a few other groups and species.

The murein sacculus functions as the structural framework of the cell wall, upon and within which various other substances are deposited and embedded. Gram-positive and Gram-negative bacteria differ both in the architecture of this framework and in The chemical composition of other cell wall constituents.

Cell wall of Gram-positive bacteria. In Gram-positive bacteria, the murein network accounts for 30–70% of the dry weight of the cell wall (which may be up to 40 layers thick). Instead of meso-diaminopimelic acid, LL-diaminopimelic acid or lysine is frequently present. In Staphylococcus aureus, the tetrapeptide side chains of muramic acid are interconnected by interpeptide bridges (e.g., pentaglycine chains). The amino acids involved in forming these structures vary from species to species, making the species-specific architecture of the supporting skeleton a reliable taxonomic marker. In the cell walls of Gram-positive bacteria, polysaccharides, if present at all, are covalently linked to the framework, while protein content is low. A characteristic feature is the presence of Teichoic Acids — chains consisting of 8 to 50 glycerol or ribitol residues linked by phosphate bridges. Some teichoic acids also contain erythritol or mannitol. Teichoic acids are presumably attached to murein via phosphodiester bonds involving phosphate groups. Cell wall of Gram-negative bacteria. In Gram-negative bacteria, the murein layer is single-layered (Fig. 2.25) and makes up less than 10% of the cell wall dry weight (in Escherichia coli). Murein contains exclusively meso-diaminopimelic acid and lacks lysine; interpeptide bridges are absent. The structure of the murein sacculus is uniform across all Gram-negative bacteria. Alongside this supporting skeleton, large quantities of Lipoproteins, lipopolysaccharides, and other lipids are anchored to the outer surface of the murein framework. These components are Covalently bound and account for up to 80% of the cell wall dry weight. Calcium Ions appear essential for maintaining the Stability of the lipopolysaccharide layer. In many Gram-negative bacteria, the murein layer becomes accessible to the murein-lysing enzyme lysozyme only after treatment with EDTA (ethylenediaminetetraacetic acid) to remove Ca²⁺. This chelating agent causes the release of a portion of the lipopolysaccharides. Teichoic acids have not been detected in Gram-negative bacteria to date.

Action of lysozyme and penicillin. The structure of the cell wall and murein was largely elucidated through investigations into the effects of lysozyme and penicillin on bacteria. Discovered by A. Fleming in 1922, lysozyme is a bactericidal enzyme found in tears, nasal mucus, and egg white, and has also been isolated from bacteria (Escherichia coli, Streptomyces) and bacteriophages. When a suspension of Gram-positive bacteria is treated with lysozyme, rapid clearing of the suspension is observed. Micrococcus luteus (lysodeikticus) undergoes lysis at a lysozyme concentration as low as 1 µg/mL. Lysis of Bacillus megaterium cells requires a concentration of 50 µg/mL, whereas many Gram-negative bacteria dissolve only when a chelating agent (EDTA) is added to the suspension.

Lysozyme cleaves the glycosidic bond in murein between carbon atom 1 of N-acetylmuramic acid and carbon atom 4 of N-acetylglucosamine. Consequently, the polysaccharide chains are broken down into disaccharide fragments composed of these two sugars (Fig. 2.25). Thus, lysozyme functions as an N-acetylmuramidase.

Complete destruction of bacterial walls can be avoided by conducting lysis in an isotonic or mildly hypertonic (0.1–0.2 M) sucrose solution. Under these conditions, lysozyme converts the cells into spherical "protoplasts" that are extremely sensitive to osmotic conditions. Protoplasts remain stable in hypertonic and isotonic media, but burst in hypotonic media, leaving behind only "ghosts" (remnants of Plasma Membranes). The term protoplasts should be restricted exclusively to rounded cells that completely lack cell wall remnants—meaning neither muramic acid nor the signature cell wall amino acid diaminopimelic acid (absent in proteins) can be detected. Lysis of the cell wall does not disrupt metabolism; protoplasts respire similarly to intact cells, form spores if sporulation was already initiated, but fail to adsorb phages.

Alongside lysozyme, numerous Other Enzymes can lyse the murein framework. Muroendopeptidases, most commonly derived from bacteria, cleave the cross-linking peptide bonds with high Specificity. For instance, an endopeptidase isolated from E. coli cleaves the bond between D-alanine and meso-diaminopimelic acid (indicated in Fig. 2.25). Other enzymes cleave bonds at different sites.

The antibiotic penicillin acts primarily on Gram-positive bacteria (staphylococci and pneumococci) as well as certain Gram-negative cells (gonococci, meningococci, enterobacteria), killing them. However, its bactericidal effect is restricted to actively growing bacteria; non-growing, "resting" cells remain unaffected. The most remarkable phenomenon observed upon penicillin treatment is the appearance of so-called L-forms, which arise from normal bacterial cells undergoing unbalanced longitudinal and radial growth. As a result, the original rod-shaped cells expand manifold in volume (see Section 3.19). On Agar nutrient media, such giant cells can remain viable for some time. When growing cells are treated with penicillin in a hypotonic solution, they lyse. In iso- and hypertonic media, the rods transform into spherical structures (Fig. 2.26) designated as L-forms or "spheroplasts." The latter differ from protoplasts in that they retain cell wall remnants (chemical analysis reveals traces of muramic acid and diaminopimelic acid). Penicillin disrupts The process of Cell wall formation.

Cell wall formation. The Biosynthesis and incorporation of murein structural units into the peptidoglycan skeleton can be divided into three stages (Fig. 2.27). The Initial Stages of biosynthesis take place in the cytoplasm, where the muramic acid pentapeptide is synthesized. The process begins with the formation of N-acetylglucosamine-1-phosphate, followed by a sequence of enzymatic reactions yielding a lactyl ether to which Five amino acids are sequentially attached. Throughout this process, the growing molecule remains linked to a uridine diphosphate (UDP) carrier. The attachment of the muramic acid pentapeptide to N-acetylglucosamine and the addition of five Glycine residues occur during the Second Stage of synthesis at the plasma membrane. For this step to proceed, the hydrophilic molecule must be converted into a lipophilic one, achieved by replacing UDP with the C₅₅-polyisoprenoid undecaprenyl phosphate. This compound facilitates The transport of the completed cell wall precursor across the plasma membrane. The Third Stage involves its incorporation into the peptidoglycan skeleton and the formation of peptide bonds. This cross-linking is accomplished via transpeptidation, in which the bond between two D-alanine residues is cleaved, and the newly freed carboxyl group forms a peptide bond with the amino group of the lysine

of the second oligopeptide, releasing the terminal D-alanine. This incorporation step also releases undecaprenyl diphosphate, which undergoes hydrolysis, and the resulting undecaprenyl monophosphate is recycled into the next cycle. This same lipid carrier also serves to transport precursors during the Synthesis of Other polymers located outside the cytoplasmic membrane, such as polysaccharides, lipopolysaccharides, and cellulose.

Fig. 2.26. Escherichia coli after treatment with penicillin (100 units/mL of nutrient medium, 90 min). 9000×. (Photo by N. Frank.)

Fig. 2.27. Biosynthesis of murein in Staphylococcus aureus. The three stages of synthesis are separated by dashed lines. They take place in the cytoplasm, at the plasma membrane, and within the cell wall. G = N-acetylglucosamine; M = N-acetylmuramic acid; UDP = undecaprenyl. Cell wall synthesis is arrested if (A) a diamino acid (lysine) is absent, (B) the racemization of L-alanine to D-alanine and/or its incorporation into the peptide are inhibited by C-cycloserine (oxamycin), or (C) penicillin prevents the cross-linking of peptide chains.

Penicillin does not affect the synthesis of the Structural components of murein, but it prevents the cross-linking mediated by transpeptidation; however, it does not interfere with the elongation of heteropolymer chains via transglycosylation. As a result of penicillin exposure, along with UDP-pentapeptide, several bacteria also release uncross-linked peptidoglycan chains into the medium. Cell wall formation can likewise be disrupted by penicillin derivatives; Cephalosporins, ristocetins, vancomycin, bacitracin, and cycloserine act in a similar manner. Spheroplasts also form in the presence of glycine and D-amino acids or as a consequence of so-called anaerobic lysis.

Outer layers of the cell walls in gram-negative bacteria.

In these bacteria, the outer layer of the cell wall lies on top of the single-layered or at most double-layered murein sacculus. Under ultrathin sectioning, it resembles the plasma membrane in appearance—this is the so-called outer membrane. This cell wall layer consists of proteins, phospholipids, and lipopolysaccharides (Fig. 2.28).

Lipoproteins are covalently bound to the murein layers—apparently via diaminopimelic acid; they are oriented with their lipophilic ends facing outward and are thus anchored within the lipophilic bilayer (via hydrophobic interactions). This layer accommodates phospholipids and the hydrophobic ends of lipopolysaccharides, whereas the hydrophilic ends of the latter face outward.

Lipopolysaccharides have gained major importance in bacteriological Diagnostics and epidemic tracking. Different strains of Salmonella typhimurium, Shigella dysenteriae, and other enteric pathogens differ from one another by the so-called O-specific side chains of lipopolysaccharides, which form the outer layer of the cell wall. Minor differences in the composition of this layer can be identified using immunological methods. Based on serological reactions within the genus Salmonella, more than a thousand species and strains have been distinguished. Chemical analysis has confirmed the differences revealed by serological methods (Westphal). There are endemic strains of Salmonella that can be identified by their immunochemical characteristics. This frequently makes it possible to determine where a patient became infected or where an epidemic originated; for instance, one can ascertain whether a given patient contracted an intestinal infection in a South American or an East Asian city.

Bacterial strains isolated from a natural source or from a patient typically grow on agar in the form of smooth, shiny colonies (S-forms, from English "smooth"); their surface contains a high amount of water due to the presence of O-specific polysaccharide chains. These S-forms spontaneously convert into R-forms (from English "rough"), which generally form rough, flat colonies (Fig. 2.29). Evidently, the polysaccharide chains confer a certain selective advantage upon the bacteria. For instance, within the host Organism, the bacteria may resist phagocytic leukocytes and are therefore extremely virulent. Only The production of Antibodies and the binding of these antibodies by bacterial polysaccharides create the conditions necessary for bacterial destruction. The immense diversity of O-specific polysaccharides in pathogenic bacteria can be explained by the Selection of continuously emerging mutant types of O-Antigens; new types confer an advantage because the host cannot possess antibodies against hundreds of antigens simultaneously. Lipopolysaccharides rank among the most potent bacterial endotoxins, inducing fever and diarrhea in patients.

Fig. 2.28. Structural model of the cell wall in gram-negative bacteria. The murein layer borders directly on the cytoplasmic membrane; the hydrophilic ends of lipoprotein molecules are covalently linked to it. The latter are embedded by their lipophilic ends into the lipid bilayer, which contains phospholipids and lipid A as a component of lipopolysaccharides. The hydrophilic O-specific heteropolysaccharide side chains of the polysaccharides are directed outward (upward in the diagram). The lipopolysaccharide molecule is shown on the right. Glu = glucose; Glu-N = glucosamine; NA = N-acetylglucosamine; Gal = galactose; Hep = heptose; KDO = 2-keto-3-deoxyoctonic acid; M = murein; OM = outer membrane; PM = plasma membrane; PP = periplasmic space.

Fig. 2.29. Colony forms of the bacterium Salmonella paratyphi B. A - S-form; B - R-form. (Photo by E. Kröger.)

The lipopolysaccharides of Salmonella typhimurium and other enterobacteria have been studied in considerable detail. Three regions should be distinguished in a lipopolysaccharide molecule: lipid A, the core region, and the O-specific side chain (Fig. 2.28). Lipid A comprises a glucosamine disaccharide whose hydroxyl groups are ester-linked to Fatty acids; this part of the molecule exhibits hydrophobic properties. Further outward lies the R-core region—a trisaccharide consisting of three residues of 2-keto-3-deoxyoctonic acid (KDO) and also bound to phosphoethanolamine; this is followed by two heptose molecules and the outer core region. The latter consists of a branched chain containing glucose, galactose, and N-acetylglucosamine. This core structure is identical in all salmonellae. In R-mutants, the molecule is shorter, terminating at the KDO trisaccharide; mutants lacking KDO have never been observed and are presumably nonviable. Adjoining the core region are the O-specific side chains; these are long chains of repeating Oligosaccharides that may contain galactose, mannose, rhamnose, abequose, fucose, colitose, and other sugars in a sequence that varies from strain to strain. The reducing C1-ends of the sugars are oriented inward. The outer heteropolysaccharide chain is strain-specific and represents the (somatic) O-antigens (Section 2.2.5). It allows the strain to be identified using the immunochemical methods mentioned above.

Function of the outer membrane. The outer membrane of gram-negative bacteria performs not only mechanical but also important physiological functions. Embedded within its lipid bilayer—which consists of lipid A, polysaccharides, and phospholipids—are proteins that span this layer entirely. Presumably, these transmembrane proteins form water-filled channels, serving as hydrophilic pores within the lipophilic membrane; hence, they are called porins. Several distinct porins exist. They allow hydrophilic low-molecular-weight substances (up to a molecular mass of approximately 6000) to pass through the membrane.

The outer membrane closely adheres to the murein layer and is linked to it by lipoproteins. The murein layer appears to be freely permeable to various substances. The space between the murein and the plasma membrane is termed the periplasmic space. It houses proteins, including depolimerases (proteinases, nucleases, such as the restriction enzyme EcoRI), peripheral Proteins of the plasma membrane, and so-called binding proteins. The latter participate in the transport of certain substrates into the cytoplasm and act as receptors for chemotactic stimuli. The periplasmic space most likely also plays a role in osmoregulation.

2.2.4 Capsules and Slime

The cell walls of many bacteria are externally enveloped by more or less thick layers of highly hydrated material with a high water content—namely, capsules and slime. Such coverings are not vital for bacteria, but possessing a capsule renders certain pathogenic bacteria resistant to phagocytosis, thereby increasing their virulence in experimental animals.

Capsules. Capsules can be visualized under a Light Microscope by adding stains such as nigrosin, Congo red, or India ink to the preparation, as these Dyes do not penetrate the capsule. This results in negative staining, where the bright capsule stands out against a dark Background (Figs. 2.30 and 2.31). Thinner pneumococcal capsules become visible upon the addition of a homologous antiserum, which triggers the deposition of antibody protein. The resulting picture resembles swelling (the Neufeld "Quellung reaction"). In most cases, the capsule consists of polysaccharides (in Streptococcus mutans, S. salivarius, Xanthomonas, and corynebacteria). In addition to glucose, capsular polysaccharides contain aminosugars, rhamnose, 2-keto-3-deoxygalactonic acid, uronic acids, and organic acids such as pyruvic and acetic acid. The capsules of certain Bacillus species (B. anthracis, B. subtilis) are composed of Polypeptides, primarily Polyglutamic acid.

Slime. Many capsular components are released into the surrounding environment as slime. Sometimes, capsules can be stripped from the cell surface by shaking or homogenizing the bacterial suspension, allowing the slime to be extracted from the nutrient medium. Particularly abundant slime production is observed in many microorganisms when the medium contains sucrose. A well-known example is the bacterium Leuconostoc mesenteroides (a representative of Heterofermentative lactic acid bacteria), which rapidly converts a cane sugar solution into a dextran jelly, earning it the name "frogspawn bacterium" in sugar refineries. This conversion occurs extracellularly and is catalyzed by an extracellular hexosyltransferase, dextransucrase:

n Sucrose + (1,6-α-glucosyl)_m → n Fructose + (1,6-α-glucosyl)_m+n

Fig. 2.30. Bacterial capsules: the purple sulfur bacterium Amoebobacter roseus (A) and the nitrogen-fixing bacterium Azotobacter chroococcus (B). Cells suspended in India ink. (A - 1200×, photo by N. Pfennig; B - 500×, photo by D. Claus.)

Dextran is a polysaccharide composed of α-D-glucose residues linked at the 1,6 position; in other words, it is a 1,6-α-glucan. Parallel polysaccharide chains are cross-linked into a single network. Dextran is a component of Blood Plasma expanders, is used to increase the viscosity of aqueous solutions, and serves as the matrix for dextran gel (Sephadex).

Fig. 2.31. Bacterial capsules. Chains of Bacillus megaterium cells suspended in India ink. Against the dark background of the ink particles, the capsules stand out as bright zones. Phase contrast, 1000×. (Photo by G. Bohlken, Diss., Göttingen, 1965.)

Cariogenic streptococci (tooth-decay-causing bacteria), including Streptococcus mutans and S. salivarius, secrete a different hexosyltransferase that converts sucrose into polyfructose (levans). These polysaccharides deposit on the tooth surface, acting as a matrix that accumulates acidic Fermentation products generated by streptococci, predominantly lactic acid.

Sheaths. Sheaths are tubular envelopes found in filamentous bacteria known as chlamydiobacteria (Sphaerotilus natans and Leptothrix ochracea). They consist of a heteropolysaccharide containing glucose, glucuronic acid, galactose, and fucose.

The targeted secretion of slime in the form of a stalk provides certain bacteria (such as Gallionella ferruginea) with limited mobility; slime can also bind individual cells into long ribbons (Zoogloea ramigera, Fig. 2.32) or leathery films (Bacteriogloea). Acetobacter aceti subsp. xylinum secretes cellulose, which aggregates cells into a tough leathery film ("Mycoderma aceti").

Cells of Sarcina ventriculi (Fig. 2.33) and Lampropedia hyalina are cemented together by cellulose into regularly shaped aggregates. In these cases, cellulose serves as a binding substance, differing in Structure and function from capsule materials. The loss of The ability to synthesize cellulose due to mutation does not affect the growth capacity of these microorganisms.

Fig. 2.32. Structure of Zoogloea ramigera colonies. A - typical colony morphology; B - individual cells embedded in the mucous ground substance. (Unz R. F., Dondero N. C., Canad. J. Microbiol., 13 [1967] 1671.)

Fig. 2.33. Cell packets of Sarcina ventriculi grown in a nutrient medium. (Bright-field micrograph, 750×; photo by D. Claus.)

Biosynthesis. Polysaccharides located exterior to the cell wall are referred to as exopolysaccharides. Based on their physical properties, a distinction is made between capsules and slime. When exopolysaccharides are relatively firmly attached to the cell wall, they are called capsules; if the attachment is loose or absent altogether, they are termed slime.

Regarding The biosynthesis of exopolysaccharides, two key points should be noted: 1) dextrans and levans are synthesized from Disaccharides by extracellular enzymes; 2) the composition of most exopolysaccharides is independent of the substrate used—their biosynthetic pathway closely mirrors the assembly mechanism of murein or lipopolysaccharides. All three processes involve UTP and the lipid undecaprenyl diphosphate (C₅₅-polyisoprenoid). Sugars activated by uridine triphosphate (UTP-sugars) attach to the carrier lipid, combine to form species-specific homo- or heteropolymeric structural components of the future polysaccharide, and are translocated from the protoplast to the outer layers of the cell wall, where they are assembled into macromolecular exopolysaccharide.

2.2.5 Flagella and Motility

Bacterial motility can be achieved through various mechanisms. In the majority of actively moving, swimming bacteria, locomotion is driven by the rotation of flagella. Flagella-free movement is characteristic of gliding bacteria (which include myxobacteria, cyanobacteria, and several other groups) as well as spirochetes. The mechanisms of their movement will be discussed when examining the respective bacterial groups.

Fig. 2.34. MAIN TYPES OF flagellation and bacterial motility patterns.

Flagellar arrangement. The arrangement of flagella in motile eubacteria is a trait characteristic of specific groups and therefore holds taxonomic significance. In rod-shaped bacteria, flagella may be inserted polarly or laterally (Fig. 2.34). Among bacteria with monopolar flagellation, only a few possess a single, exceptionally thick flagellum—these are monotrichous (Vibrio metschnikovii, Fig. 2.35; Caulobacter sp.). In many bacteria with monopolar and bipolar flagellation, what appears to be a single flagellum is actually a bundle of 2 to 50 flagella (polytrichous). Monopolar polytrichous flagellation is also referred to as lophotrichous (as in Pseudomonas, Chromatium), whereas bipolar polytrichous flagellation is termed amphitrichous (in Spirillum). Selenomonas features a single bundle of flagella inserted laterally (Fig. 2.36, B). In peritrichous arrangement (as in Enterobacteriaceae, Bacillaceae, and certain other bacteria), flagella are distributed laterally or across the entire cell surface (Fig. 2.36, A).

Fig. 2.35. Types of flagellation (electron micrographs). A. Aquaspirillum serpens with a polarly inserted flagellar tuft (peritrichous); 11,000×. B. Vibrio metschnikovii with a polarly inserted single flagellum (monotrichous); 7,100×. (Metal shadowing; photo by W. van Iterson.)

Fig. 2.36. Types of flagellation (electron micrographs). A. Proteus mirabilis with peritrichous flagellar arrangement (platinum-palladium oblique shadowing, 9,500×; photo by H. Frank). B. Selenomonas ruminantium with lateral flagella (negative staining, 4,080×; photo by V. Kingsley.)

Fig. 2.37. Thiospirillum jenense. (Phase contrast, 1,200×; photo by N. Pfennig.)

Visualization of flagella. Observing a flagellum (or flagellar tuft) under transmitted light or Phase-contrast microscopy is possible in only a few bacteria, such as Chromatium okenii, Bdellovibrio, and Thiospirillum (Fig. 2.37). In many other bacteria (Pseudomonas, Spirillum, etc.), flagella and their beating zone can only be visualized using dark-field microscopy. Flagella are most easily detected by staining or metal deposition techniques, as well as by electron microscopy.

Functions of flagella. In most polar-flagellated bacteria, flagella function similarly to a ship's propeller, pushing the cell through the surrounding liquid medium. The flagellum is a helically curved filament driven into rotational motion by a "motor" located at its insertion site in the plasma membrane. Cell locomotion can be driven by a single flagellum or a bundle of flagella. Flagella rotate relatively rapidly; for instance, in spirilla, they perform about 3,000 revolutions per minute, which is close to the speed of an average electric motor. The Rotation of the flagella causes the cell body to rotate in the opposite direction at approximately one-third of that speed.

Flagella can spontaneously or in response to an external stimulus reverse their direction of rotation (Fig. 2.34). In some polar-flagellated bacteria, this causes the cell to reverse its swimming direction. When Chromatium okenii reverses its flagellar rotation in response to a light flash, the flagellar tuft transforms into a pulling apparatus, causing the cell to move backward four times slower than forward, resulting in a "tumbling" motion. In Thiospirillum jenense—a giant phototrophic spirillum—the single polar flagellar tuft during reverse movement no longer beats in front of the cell; instead, the flagellar beating envelope now surrounds the sides of the cell, effectively turning itself inside out (much like an umbrella inverted by the wind). In spirilla with amphitrichous flagellation, depending on the circumstances, either one or the other tuft assumes this configuration.

Peritrichously arranged flagella of Escherichia coli operate as a single well-coordinated helical bundle, propelling the cell through the medium. Whenever the rotation direction of individual flagella reverses, the cell begins to "tumble." Apparently, peritrichously arranged flagella cannot function as a pulling apparatus.

Flagellated bacteria can move very rapidly: Bacillus megaterium at a speed of 1.6 mm/min, and Vibrio cholerae at 12 mm/min. This corresponds to roughly 300 to 3,000 body lengths per minute.

Fine structure of flagella. Flagella are helically coiled filaments. In different bacteria, they vary in thickness (12–18 nm), length (up to 20 µm), as well as in pitch and amplitude. These parameters are characteristic of each species. Some bacteria may form flagella of different types. Flagellar filaments consist of a specific protein, flagellin. They are built of subunits with a relatively low molecular mass. The subunits are arranged helically around a central hollow space (similar to the protein molecules in tobacco mosaic virus; see Section 4.1). Thus, the structure of the flagellum is determined by The properties of its protein subunits.

A flagellum consists of three parts: the helical filament described above, a "hook" near the cell surface, and a basal body. By means of the basal body, the flagellum is anchored in the cytoplasmic membrane and the cell wall (Fig. 2.38). It consists of a central rod bearing two pairs of rings in Gram-negative bacteria. The outer pair (L and P rings) is located at the level of the outer and inner layers of the cell wall, while the inner pair (S and M rings) is located at the level of the outer layer of the cytoplasmic membrane. Since Gram-positive bacteria lack the outer pair of rings, it is assumed that only the inner pair is required for flagellar rotation. One can imagine that the M ring acts as a drive disk, whereas the S ring plays The Role of a bearing on the inner surface of the peptidoglycan layer. The Molecular Mechanism of the rotational "motor" of the flagellum has not yet been elucidated.

Fig. 2.38. Attachment of the flagellum to the cell wall and cytoplasmic membrane in a Gram-negative bacterium. H – flagellar filament; Kp – hook; PM – cytoplasmic membrane; LS – lipopolysaccharide layer; PG – peptidoglycan layer; Ст – rod.

O and H antigens. Proteus vulgaris often spreads across the entire surface of agar as a thin gray film (H-form, from the German Hauch, meaning film or haze). This "swarming" is explained by the high motility of the cells. Some strains do not form a film (O-form, from the German ohne Hauch, meaning without film). These strains are non-motile and lack flagella. This is the Origin of the conventional terminology adopted in bacterial serodiagnosis: surface or general cell body (somatic) antigens are called O antigens, and flagellar antigens are called H antigens.

Fimbriae and pili. The surface of certain bacteria is covered with A large number (ranging from 10 to several thousand) of long, thin, straight filaments 3–25 nm thick and up to 12 µm long, called fimbriae or pili. They occur in both flagellated and non-flagellated species. They should be distinguished from sex pili, or F-type pili, which were discovered on donor cells of Escherichia coli K12, i.e., strains carrying the sex factor F (F⁺, Hfr). F pili occur only one or two per cell and appear as hollow protein tubes from 0.5 to 10 µm in length.

Chemotaxis. Free-swimming bacteria are capable of taxis—directed movements determined by external stimuli. Depending on the environmental factors causing the directed movement, one speaks of chemotaxis, aerotaxis, phototaxis, and magnetotaxis.

Motile bacteria react to chemical stimuli by accumulating in certain areas and avoiding others. This response of free-swimming organisms is called chemotaxis. Bacterial accumulations are formed under The Influence of Chemical factors in the following manner (Fig. 2.39). In forms with peritrichous flagella, only Two Types of locomotor behavior are possible: smooth swimming and tumbling. The latter interrupts the smooth run and alters the direction of travel. When a bacterium enters an environment with a concentration gradient of a substrate that "attracts" it (an attractant), its smooth swimming lasts for many seconds if it swims toward the optimal concentration; however, such movement ceases within a few seconds if the bacterium swims in the opposite direction. Although the direction of smooth swimming after a tumble is entirely random, the dependence of the duration of such movement on its direction ultimately results in the accumulation of bacteria in the region of optimal Substrate Concentration. Chemoreceptors are responsible for sensing and responding to the chemical stimulus. In some cases, these chemoreceptors operate independently of the bacteria's ability to utilize the given substrate. For example, some mutants continue to respond perfectly normally to a specific nutrient even though they have lost the ability to use it.

Fig. 2.39. Chemotactic movements. A. In a normal (isotropic) environment, a bacterium swims almost in a straight line and tumbles at approximately regular intervals. B. In a concentration gradient of an "attracting" substance (attractant), the tumbling frequency decreases when the cell swims in the "correct" direction, i.e., toward the concentration optimum. C. In a gradient of a repelling substance (repellent), the cell behaves in the opposite manner.

Aerotaxis. In motile bacteria, the type of metabolism (aerobic or anaerobic) can be determined by their aerotactic movements and accumulation at specific distances from the edge of the coverslip. In a layer of bacteria placed between a microscope slide and coverslip, aerophilic bacteria accumulate at the edge of the coverslip or in the immediate vicinity of air bubbles trapped in the preparation; this indicates their requirement for aerobic conditions and that they obtain the necessary energy through respiration (Fig. 2.40). Strictly anaerobic bacteria accumulate in the center. Microaerophilic bacteria, such as certain pseudomonads and spirilla, maintain a certain distance from the edge. Using bacteria that exhibit positive aerotaxis, Engelmann successfully demonstrated oxygen evolution by locally illuminated chloroplasts of the green alga Spirogyra.

Fig. 2.40. "Respiration figures" (after Beijerinck) as a consequence of aerotaxis in motile bacteria. Aerobic bacteria accumulate at the edge of the coverslip and around air bubbles trapped in the medium (A). Microaerophilic bacteria remain at some distance from the edges of the coverslip (B). Strictly anaerobic bacteria accumulate in the center (C).

Phototaxis. Phototrophic purple bacteria require light to obtain energy. It is therefore not surprising that, as a result of phototaxis, they accumulate in illuminated areas. If a preparation in which a dense suspension of Chromatium cells is uniformly distributed under a coverslip is kept in the dark and then exposed to a focused beam of light, the bacteria concentrate within the illuminated spot. Cells that accidentally enter this spot as a result of their random movement can no longer leave it. As soon as they enter the dark zone, the direction of flagellar movement instantly reverses, and the cells return to the illuminated area. The change in flagellar activity occurs so rapidly that this response has been termed the "shock reaction" (phobotaxis). Indeed, even a slight difference in illumination between two areas is sufficient to trigger this response. Small Chromatium cells accumulate in places where the illumination is only 0.7% higher than in the surrounding area. Thus, in their sensitivity to light contrast, they approach the retina of the human eye (for which the corresponding threshold is 0.4%).

Magnetotaxis. Bacteria (rods, spirilla, cocci) capable of orienting themselves in a magnetic field and moving along magnetic field lines have been isolated from the surface layers of bottom sludge in freshwater reservoirs and seas. They contain a large amount of iron (0.4% of dry weight) in the form of ferromagnetic iron oxide (magnetite), which is located in granules (magnetosomes) situated near the attachment sites of the flagella. Bacteria isolated in the Northern Hemisphere "seek" the north; here, the magnetic field lines run downward at an angle of about 70° to the horizontal, deep into the water body. Magnetotactic behavior directs bacteria into the depths of the sludge, where oxygen is very scarce or absent entirely. Since magnetotactic bacteria are anaerobes or microaerophiles, their response to the magnetic field makes ecological sense. Such cells introduced into the Southern Hemisphere will, of course, perish en masse; only a few "incorrectly" polarized cells will survive and subsequently multiply. The polarity is evidently not genetically fixed.

2.2.6 Reserve substances and other intracellular inclusions

In many microorganisms under specific environmental conditions, substances that can be regarded as reserves—polysaccharides, lipids, polyphosphates, and sulfur—are deposited inside the cells. These substances accumulate when the nutrient medium contains the corresponding starting compounds, while bacterial growth is restricted or entirely impossible due to the lack of certain nutritional components or the presence of inhibitors. Reserve substances are contained within cells in an osmotically inert form—they are insoluble in water. Under conditions favorable for growth, when these substances are needed, they are re-entered into metabolism. Reserve Polysaccharides, neutral lipids, and poly-β-hydroxybutyric acid can serve as sources of both energy and carbon. Therefore, in the absence of external Energy Sources, they can prolong cell survival, and in spore-forming species, they can provide conditions for spore formation even in the absence of exogenous substrates. Polyphosphates can be viewed as a reserve source of phosphate, and reserve sulfur as a potential electron donor.

Polysaccharides. Reserve CARBOHYDRATES of microorganisms are still poorly understood. In some microbes, starch (blue coloration) or glycogen (brown coloration) can be identified using a color reaction with Lugol's iodine solution. Unlike cell wall polysaccharides, reserve polysaccharides are all formed from a-D-glucose; the glucose molecules are linked together by 1,4-a-glucosidic bonds and extensively cross-linked (see also Fig. 14.2).

Due to a-glucosidic bonds, polyglucose chains are not elongated, but helically coiled. Starch, which plants deposit in the form of starch granules, consists of amylose and amylopectin. The proportion of amylose is 20–30% (by weight). It is this component that is responsible for the blue coloration with iodine; this reaction is based on the incorporation of iodine into the turns of the helix (inclusion reaction).

A starch-like compound in clostridia has been named "granulose" or "iodogen". Cells of Clostridium butyricum are almost entirely filled with small granules of this substance; only the pole of the cell where the spore is formed remains free of them. Cells of Acetobacter pasteurianus and many Neisseria species contain starch.

Glycogen, also called "animal starch," is similar to amylopectin, but its chains are even more branched (through the formation of 1,6-bonds); in bacteria, it appears to occur more frequently than starch. Glycogen has also been detected in Yeasts and other fungi, in bacilli (Bacillus polymyxa), in Salmonella, Escherichia coli (Fig. 2.41) and other Enterobacteriaceae, in Micrococcus luteus, and in Arthrobacter.

Fig. 2.41. Glycogen Inclusions in Escherichia coli B cells. Glycogen accumulation occurred during 15 hours of cell aeration in phosphate buffer containing 0.5% glucose. Transmission electron micrograph of an ultrathin section (osmium tetroxide-dichromate; uranyl acetate; 8,000×). (Photo by R. Hillmer and F. Amelunxen.)

Fig. 2.42. Accumulation of poly-β-hydroxybutyric acid in cells of Alcaligenes eutrophus. The cells are taken from a culture in the exponential growth phase (A), and then after 1 h (B) and 24 h (C) of acetic acid assimilation in the absence of a nitrogen source. Transmission electron micrographs of ultrathin sections (osmium tetroxide-dichromate; uranyl acetate-lead citrate; 6,000×). (Photos by R. Hillmer and F. Amelunxen.)

Fat-like substances. Granules and droplets of fat frequently occur as inclusions within microbial cells. They are visible under a light microscope due to their strong light-refracting properties and can also be stained with lipophilic dyes such as Sudan III or Sudan Black B.

The "sudanophilic" granules of many bacteria consist of poly-β-hydroxybutyric acid, a polyester that is soluble in chloroform, insoluble in ether, and composed of approximately 60 β-hydroxybutyrate residues. The proportion of this substance can reach up to 80% of the dry biomass. Poly-β-hydroxybutyric acid is synthesized by many aerobic bacteria (Fig. 2.42), as well as by cyanobacteria and anaerobic phototrophic bacteria (Fig. 2.43, A). In facultative anaerobes and strict aerobes, it accumulates when cells experience an O₂ shortage and switch to fermentation; therefore, it can be regarded as a polymeric intracellular fermentation product. Under aerobic conditions, this product can re-enter metabolism as an energy and carbon source and be utilized via respiration.

Fig. 2.43. Cells containing poly-β-hydroxybutyric acid inclusions. A — Chromatium okenii; B — Bacillus megaterium (negative stain in India ink). The dark inclusions within the C. okenii cells are sulfur droplets. Bright-field micrographs; 1,000×. (Schlegel H.G., Arch. Microbiol., 42 [1962]; Bohlken G., Diss., Göttingen, 1965.)

Neutral fats (triglycerides), which are deposited in particularly large amounts within the vacuoles of yeasts and other fungi, closely resemble the fats of higher organisms. Yeasts (Candida, Rhodotorula) can accumulate up to 80% of their dry weight in lipids.

Mycobacteria, Nocardia, and actinomycetes accumulate different lipid-like substances within their cells and often even secrete them into the medium. Mycobacteria may contain up to 40% Waxes (esters of long-chain Fatty Acids and alcohols).

The accumulation of reserve fats is determined by the COMPOSITION OF THE nutrient medium (a high C/N ratio), and these fats can be extracted directly from the cells. The amounts of other lipid components are largely independent of the medium composition. These lipids are released only after the Hydrolysis of Proteins and polysaccharides and represent components of lipoproteins—which form part of the plasma membrane and internal membranes—as well as lipopolysaccharides.

Polyphosphates. Many bacteria and green algae are capable of accumulating phosphoric acid in the form of polyphosphate granules. Such granules were first described in Spirillum volutans, which is why they are referred to as volutin granules; another term for them, "metachromatic granules," stems from their ability to induce a characteristic color change (metachromasia) in certain dyes (such as methylene blue and toluidine blue). These granules consist mainly of long-chain polyphosphates similar to Graham's salt (sodium metaphosphate); cyclic metaphosphates frequently found in analyses are likely artifacts resulting from The breakdown of polyphosphates.

Volutin granules act as phosphate reserves, enabling the cell to undergo a few additional divisions if phosphorus becomes scarce in the environment. Polyphosphates play only a minor role as energy sources.

Sulfur. In many bacteria that oxidize sulfide to sulfate, sulfur is temporarily stored as highly refractive globules. The sulfur accumulated inside cells, much like that excreted externally, exists in a liquid state and gradually transitions into its orthorhombic modification. The amount of sulfur stored depends on the hydrogen sulfide concentration in the environment: in the absence of H₂S, sulfur is oxidized to sulfate. For aerobic sulfur bacteria (Beggiatoa, Thiothrix, Achromatium, Thiovulum; Fig. 2.44), sulfur serves as an energy source, whereas for anaerobic phototrophic purple sulfur bacteria (Chromatium), it acts as an electron donor. Sulfur inclusions, occasionally found in cyanobacteria and Sphaerotilus natans, can be viewed as products of detoxifying hydrogen sulfide, which is frequently present in the habitats of these organisms.

Fig. 2.44. Colorless hydrogen sulfide-oxidizing bacteria. A. Beggiatoa gigantea with sulfur inclusions (bright-field micrograph, 300x). B. Rosettes of Thiothrix (phase contrast, 300x). C. Achromatium oxaliferum with calcium carbonate and sulfur inclusions (200x). D. Thiovulum with sulfur inclusions (1000x). [Photographs by K. Schmidt (A, C), D. Claus (B), J. W. M. La Riviere and H. Schuur (D).]

Other cellular inclusions. In Bacillus thuringiensis and related species (B. laterosporus, B. medusa), crystal-like inclusions—parasporal bodies composed of protoxin—can often be observed adjacent to the spores (see Fig. 3.6). The protoxin dissolves in the gut juice of susceptible insects (caterpillars), and the released toxin destroys the intestinal epithelium, resulting in the caterpillar's death. Preparations derived from these bacilli, which are toxic only to a few insect groups, have already been successfully employed in biological pest control.

Gas vacuoles. Many aquatic bacteria, particularly phototrophic ones, as well as colorless forms (Pelonema, Peloploca), halobacteria (Halobacterium halobium), and certain clostridia, contain gas vacuoles. These structures grant cells the ability to alter their average density and remain suspended in water. Thanks to this capability, some bacteria in stratified lakes can maintain their position within a specific water layer where growth conditions are optimal, without resorting to active flagellar swimming. Anoxygenic (non-O₂-evolving) phototrophic bacteria, including purple (Lamprocystis, Amoebobacter, Thiodictyori) and green (Pelodictyon) species, thrive in the anaerobic zone (hypolimnion) of water bodies just below the thermocline (see Fig. 17.1). The buoyancy of these organisms is apparently sufficient to keep them suspended in the cold (denser) water layer of the hypolimnion, but it does not provide enough lift to keep them in the warm (lighter) water layer above the thermocline. Oxygenic (i.e., O₂-producing) cyanobacteria (Oscillatoria agardhii, Aphanizomenon flos-aquae, Microcystis aeruginosa) flourish in layers above the thermocline. Their buoyancy is presumably regulated via photosynthesis, cellular turgor, and alterations in the number and size of gas vesicles.

Each gas vacuole is an aggregate of gas vesicles. These vesicles are spindle-shaped (cylinders with conical ends). Their shell is not made of a conventional membrane, but rather of a pure protein with a folded structure, measuring a mere 2 nm in thickness. Micrographs reveal Ribs arranged along the cylindrical portion of the vesicle like hoops on a barrel. The envelope is constructed from protein subunits with a Molecular Weight of 14∙10³. The protein molecules are evidently oriented such that the inner side of the wall is hydrophobic, while the outer side is hydrophilic. The cell contains numerous gas vesicles aligned parallel to one another. Under a light microscope, such an aggregate of gas vesicles (i.e., a gas vacuole) appears as an optically empty, highly refractive region.

Carboxysomes. The cells of certain autotrophic bacteria contain carboxysomes. These are polyhedral bodies about the size of a phage HEAD, which enclose the enzyme ribulose-1,5-bisphosphate carboxylase (often alongside a small amount of DNA). Carboxysomes have been found in Nitrosomonas, Thiobacillus, and numerous cyanobacteria.

2.2.7 ENDOSPORES AND OTHER Resting Forms

Only a small group of bacteria are capable of endospore formation. The immense significance of endospores stems from their thermostability. While nearly all other bacteria, as well as the vegetative cells of spore-forming species, perish at 80°C (pasteurization temperature) within 10 minutes, heat-resistant endospores withstand significantly harsher heating; some spores even survive boiling for many hours. Labor-intensive and costly sterilization techniques are specifically designed to eradicate endospores. On the other hand, the thermostability of spores offers a unique opportunity for the selective enrichment of spore-forming cultures. Soil or another sample collected from a specific Location is heated at 80 or 100°C for 10 minutes, which destroys the vegetative cells; only the heat-resistant spores remain viable and germinate in a suitable nutrient medium.

Classification of endospore-forming bacteria. With a single exception, spore-forming species belong to rod-shaped Gram-positive bacteria. Most of them are motile by means of peritrichous flagella. Bacteria belonging to the genus Bacillus are strict aerobes or facultative anaerobes. The genera Clostridium and Desulfotomaculum comprise anaerobic, spore-forming bacteria. Clostridia obtain energy via fermentation, whereas Desulfotomaculum species can generate energy through Anaerobic respiration, utilizing sulfate as an electron acceptor. Sporolactobacillus is among the lactic acid bacteria. Sporosarcina possesses spherical cells, yet physiologically it aligns with the bacilli. The GC content (see section 22.1) of DNA in spore-forming cells is remarkably low; clostridia, whose DNA contains 22 to 27 mol % GC, sit at the very end of prokaryotes ordered by decreasing values of this metric.

Detection of endospores. Under microscopic examination, spores are visible due to their high refractive index—comparable to that of dehydrated protein—which indicates that a large amount of protein-rich material is concentrated within a small volume in spores. A spore contains nearly all the dry matter of the mother cell while occupying one-tenth of its volume. In doubtful cases, whether cells contain true endospores can be determined using specialized staining. If a heat-fixed bacterial smear is boiled with carbol fuchsin, the spores firmly bind the dye and do not decolorize even upon treatment with ethanol or 1 M acetic acid, whereas the rest of the cellular contents becomes colorless.

Sporulation. Spores are formed within the bacterial cell. This process begins with the accumulation of protein material, causing the refractive index at the sporulation site to increase. The accompanying metabolic shifts entail the consumption of reserve substances (poly-β-hydroxybutyric acid in aerobes and polysaccharides in anaerobes). During the first five hours of sporulation, a significant portion of the mother cell's proteins breaks down. This yields dipicolinic acid (pyridine-2,6-dicarboxylic acid), a substance unique to spores. This acid is absent in vegetative cells. Concurrently with dipicolinic acid synthesis, calcium ions are taken up; in mature spores, this acid presumably exists as a calcium chelate and can account for 10–15% of the spore's dry weight. Dipicolinic acid is localized within the spore protoplast and occurs exclusively in heat-resistant endospores (Figs. 2.45, 2.46).

Spore formation is one of the most complex processes of bacterial Cell Differentiation. It begins with a specialized, asymmetric cell division (Fig. 2.45). As a result of the invagination of the plasma membrane, a portion of the protoplast is pinched off from the mother cell. This protoplast contains a fraction of the nuclear material—a single genome. Unlike normal cell division, a cell wall is not formed between the two protoplasts. Instead, the protoplast of the future spore becomes surrounded, or enveloped, by the plasma membrane of the mother cell. Consequently, it is enclosed by two plasma membranes, each of which participates in the synthesis of the spore wall. The membrane of the spore protoplast synthesizes the germ cell wall on its outer side, while the membrane derived from the mother cell synthesizes the spore cortex inward. The cortex consists of a multi-layered peptidoglycan framework that differs from the wall skeleton of vegetative cells, among other things, in its degree of cross-linking. The outer coat of the spore is formed by the mother cell and largely consists of polypeptides. The mother cell also produces an additional, thin polypeptide layer known as the exosporium; this structure is present in only a few bacteria (such as Bacillus cereus) and surrounds the spore like a loose sheath. Given this multi-layered envelope, it is not surprising that the coat accounts for about half the volume and dry mass of a mature spore.

Induction of sporulation. Spores are by no means an obligatory stage in the bacilli life cycle; under favorable nutritional conditions, bacilli can multiply indefinitely by division as vegetative cells. Spore formation is initiated only when nutrients become scarce or when Metabolic waste products accumulate in excess. In other words, it occurs exclusively when promoted by environmental conditions. Desiccation does not stimulate sporulation. When vegetative cells are placed in distilled water, "endotrophic sporulation" can be observed—that is, the formation of spores at the expense of intracellular reserve materials. In such cases, spore formation is clearly triggered by a deficiency of the exogenous substrate. The induction of sporulation takes place over several hours. If, for example, glucose is added to a suspension of Bacillus cereus var. mycoides vegetative cells within the first 5 h after they are transferred to water, spore formation ceases—the addition of the substrate suppresses sporulation. Glucose added later than 6 h has little effect, and sporulation is no longer inhibited. The induction (derepression) of sporulation proceeds, and within 10–13 h after the cells are transferred to water, approximately 90% of them form spores. Therefore, sporulation is regulated by external factors.

Fig. 2.45. Schematic diagram of sporulation and the structure of a mature spore. A, B — process of spore protoplast separation; C, D, E — formation of the forespore; F — mature spore. 1 — cytoplasm; 2 — plasma membrane; 3 — germ cell wall; 4 — spore cortex; 5 — inner spore coat; 6 — outer spore coat; 7 — exosporium. (W. G. Murrell.)

Fig. 2.46. Morphological and physiological changes during spore formation in aerobic bacteria. This process is triggered by the utilization of glucose (zero time). Spore formation can be monitored by the increase in calcium and dipicolinic acid content, as well as by the growing number of heat-resistant spores and changes in light refraction. Specific biochemical shifts correspond to morphological changes. The stages I — VII are schematically depicted at the top.

The number of spore-forming cells often increases following the addition of manganese salts to the medium.

The ability to form endospores is gradually lost upon repeated subculturing of vegetative cells. Because Suspensions of spore-forming microorganisms typically contain both spores and vegetative cells, the culture is generally subjected to brief boiling before each transfer. This helps preserve or enhance the cells' capacity for sporulation.

Properties of mature spores. Spores are released through the autolysis of the mother cells. Mature spores exhibit no metabolic activity whatsoever. They are exceptionally resistant to high temperatures, various types of radiation, and chemical agents. This heat resistance is attributed to a very low water content. Spores of Bacillus megaterium contain only about 15% water—roughly the same as wool or dry casein. Lyophilized vegetative cells of bacteria are also highly heat-tolerant. The thermoresistance of spores is approximately proportional to their dipicolinic acid content.

The Radioresistance of spores is likewise higher than that of vegetative cells. It is roughly proportional to the concentration of disulfide groups in the outer protein layer. The spore coat consists primarily of a Cysteine-rich protein resembling keratin. Meanwhile, the chemical resistance of endospores is due to the impermeability of their coats to many substances.

Spore germination. In suitable media, the majority of spores germinate. Appropriate pretreatment, specific storage conditions, and heat activation can enhance spore "germination capacity"—increasing the percentage of germinated spores. For Bacillus subtilis, optimal conditions for stimulating germination are considered to be a seven-day resting period followed by a five-minute heat treatment in water at 60°C. Other spores can be activated by brief boiling (10 min at 100°C). Heat-shock treatment must be performed immediately before plating the spores, as the activation process is reversible. Germination is preceded by Water uptake and swelling. In some cases, the presence of glucose, amino acids, nucleosides, or Other Compounds is required for the germination of activated spores. During germination, profound physiological changes occur: respiration and enzymatic activity increase rapidly; the release of amino acids, dipicolinic acid, and Peptides begins. Dry matter loss during germination reaches 25–30%. As they germinate, spores lose their heat resistance. The emerging germ tube is surrounded by a very thin and apparently incompletely formed cell wall, allowing even DNA to penetrate the protoplast (see the section on transformation). The germ tube may form in either a polar or lateral position; in some instances, the spore coat ruptures, whereas in others, the germ tube pierces through it (Fig. 2.47).

Fig. 2.47. Germinating spores. 1 — polar germination of a Clostridium spore (with exosporium); 2 — polar germination of a Bacillus megaterium spore (spore coats remain attached); 3 — lateral germination of spores in bacilli (a — B. cereus; b — B. subtilis).

Longevity of spores. Bacteria can remain in a state of suspended animation (cryptobiosis) as spores for extended periods. Only a small Number of viable Bacillus subtilis and B. licheniformis spores were found in soil adhered to plant specimens from the Kew Gardens herbarium (England) that had been stored in a dry state for 200 to 320 years. Spores of B. coagulans and B. circulans were also detected in soil samples stored for 50 to 100 years. According to such experiments, up to 90% of spores lose their viability after 50 years of storage in dry soil. Judging by this, one ton of dry soil would still contain viable spores even after 1,000 years.

In the dry state, many bacteria (if not most) retain viability for a number of years. For preservation in culture collections, vegetative cells are generally subjected to freeze-drying (lyophilization) and stored at room temperature or at low temperatures under vacuum. As calculated long ago by Becquerel, microorganisms can remain viable for millions of years at temperatures close to absolute zero. Short-term experiments with liquid nitrogen and the extrapolation of their results lead to the conclusion that such assumptions are entirely well-founded. Bacteria that do not tolerate lyophilization can withstand multi-year storage in suspensions at liquid nitrogen temperatures.

Other resting forms (cysts, exospores, myxospores). Endospores are long-lived survival forms of bacteria that are resistant to high temperatures, desiccation, radiation, and chemical agents. In addition to endospores, certain other bacteria possess alternative resting forms—exospores and cysts. The formation of exospores has hitherto been observed only in the methane-metabolizing bacterium Methylosinus trichosporium¹. Exospores arise through the budding of the mother cell and are similar in properties to the endospores of bacilli. Some bacteria form spherical, thick-walled cells called cysts. Upon nutrient depletion, the entire rod-shaped vegetative cell transforms into a cyst, rather than just a part of it as in endospore formation. Cysts of Azotobacter species, as well as Methylocystis, are resistant to drying, mechanical stress, and radiation, but not to high temperatures. A similar transformation of an entire cell into a cyst underlies the formation of myxospores from rod-shaped vegetative cells in Myxococcus and Sporocytophaga.

Cells of Arthrobacter (A. globiformis) are pleomorphic. When nutrients are abundant, they grow as rods; however, when the nutrient supply is exhausted, coccoid cells appear. Arthrobacter belongs to those bacteria capable of surviving in a dormant state in dried soil for some time, with no structural differentiation having been observed in this case.

¹ Exospores are also formed by the phototrophic purple bacterium Rhodomicrobium vannielii. — Ed. note.

2.2.8 Bacterial and Fungal Pigments

Colonies of many bacteria and fungi exhibit striking coloration due to the secretion of pigmented products into the environment or the pigmentation of the cell itself. The capacity for pigment production is genetically determined and can therefore be used as a diagnostic feature. Pigmented forms are easier to detect and identify. The pigments encompass representatives of various chemical classes: carotenoids, phenazine dyes, pyrroles, azaquinones, anthocyanins, etc. (Fig. 2.48).

Protection against light and ultraviolet radiation. In Petri dishes containing complex agar media that have been left open for a time in dusty air, colored colonies of microorganisms frequently appear. Their coloration is dominated by yellow, orange, and red hues caused by the presence of carotenoids. Among the bacteria forming such colonies, we most commonly find representatives of the genera Micrococcus, Corynebacterium, Mycobacterium, and Nocardia, and among yeasts, Rhodotorula. The Abundance of pigmented forms among the "aerial microflora" is explained by the protective role of pigments, which shield cells from the effects of visible and near-ultraviolet light. Consequently, in exposed habitats (dust, straw), colorless bacteria perish faster than pigmented ones. The protective action of pigments against radiation damage can also be demonstrated using an orange, carotenoid-containing halophilic bacterium and its colorless mutants. While both the wild type and the mutants grow equally well under dim light, the growth of the colorless mutants is severely inhibited under bright sunlight. The bactericidal effect of visible light manifests only in the presence of molecular oxygen and is driven by photooxidation; certain cellular pigments (flavins and cytochromes) act as catalysts (photosensitizers) in this process. Carotenoids are located within the plasma membrane and protect vulnerable cellular regions from the effects of photooxidation.

Fig. 2.48. Selected pigments produced by bacteria and yeasts. Pyocyanin is secreted by cells of Pseudomonas aeruginosa, and iodinin by Chromobacterium iodinum. Indigoidine is produced by various bacteria, including Pseudomonas indigofera. Prodigiosin is formed by Serratia marcescens (formerly Bacterium prodigiosum), and violacein by Chromobacterium violaceum. Pulcherrimin is a pigment of the Yeast Candida pulcherrima. Flexixanthin is the main carotenoid of Mycobacterium phlei and is a carotenoid glycoside; sarcinaxanthin is the primary carotenoid of yellow Sarcina species (Micrococcus luteus).

Photosensitization. The natural sensitivity of cells to molecular oxygen, which becomes apparent in the light, can be enhanced. Bacteria stained with vital dyes—such as methylene blue, eosin, or acridine orange—and subsequently exposed to light die off faster than unstained cells. The dye molecule absorbs light and is capable of transferring its energy to an O₂ molecule. In the process, the oxygen molecule, normally in a triplet state, is elevated to an excited state (singlet oxygen). This singlet oxygen initiates oxidation reactions that ordinary oxygen cannot perform (cycloadditions, ene reactions). Photosensitization is utilized in fur farms and zoos to eliminate potentially pathogenic bacteria; for this purpose, methylene blue or other dyes are added to the drinking water. Photosensitized bacteria perish upon exposure to normal daylight.

Carotenoid biosynthesis. The intense red color of purple bacteria is due to the presence of red carotenoids (possessing 12–13 double bonds and methoxy and oxo groups). Here, the pigments play not only a protective role but also absorb light for photosynthesis and participate in light reception during phototaxis. Carotenoids, along with bacteriochlorophylls, are located in photosynthetically active membranes (thylakoids, chromatophores).

In many pigmented microorganisms, pigment formation (much like the synthesis of photosynthetic pigments in higher plants) occurs only in the presence of light. Mycobacteria, including the pathogenic tubercle bacillus (Mycobacterium tuberculosis), synthesize carotenoids only when exposed to light. The same applies to bacteria growing on cured ham or cheese. In many cases, pigmentation depends on the composition of the nutrient medium and temperature.

Pulcherrimin. The color of red yeasts (Rhodotorula, Sporobolomyces salmonicolor) is primarily determined by carotenoids. Pulcherrimin, a pigment of Candida pulcherrima, belongs to a different class of substances. Along with C. reukaufii, these yeasts can be isolated from nectar-bearing flowers and fruits, as well as from the gut of bees. On media containing iron, they form dark-red colonies. Their red pyrazine pigment, which is insoluble in water and other Solvents, contains complex-bound iron.

Prodigiosin. On media containing carbohydrates, the bacterium Serratia marcescens (formerly known as Bacterium prodigiosum) frequently develops. The bright red color of its colonies (as well as cell suspensions) is due to the presence of the pigment prodigiosin, whose molecule contains three pyrrole rings. This pigment is also found in actinomycetes.

Indigoidine. Indigoidine belongs to the azaquinones (diazadiphenoquinones) and is a water-insoluble blue pigment secreted into the medium by various bacteria, such as Pseudomonas indigofera, Corynebacterium insidiosum, Arthrobacter atrocyaneus, and A. polychromogenes.

Violacein. Chromobacterium violaceum is easily isolated from soil by placing rice grains in a Petri dish on heavily moistened soil; the colonies of this microorganism are readily identified by their bluish-purple color, caused by the water-insoluble purple pigment violacein. This pigment is an indole derivative formed during the oxidation of Tryptophan.

Phenazine pigments. Many pigments secreted into the environment by aquatic bacteria belong to phenazine derivatives. The best known among them is pyocyanin, produced by cells of Pseudomonas aeruginosa (formerly P. pyocyanea). Various strains and species of pseudomonads secrete phenazine-1-carboxylic acid, oxychlororaphin, or iodinin, and sometimes all of these pigments simultaneously.

Secondary metabolites. In many microorganisms, pigments are secondary metabolites, meaning they do not belong to the universal compounds found in all such organisms (see Section 10.4). Their structure clearly shows that they are derivatives of common metabolites or structural cell components. Some pigments exhibit antibiotic properties, making many pigmented microorganisms antibiotic producers. There is such a close correlation between pigmentation and the Formation of secondary metabolites that the presence of pigments strongly suggests the likely production of antibiotics and other BIOLOGICALLY ACTIVE SUBSTANCES.