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E. Coli metabolization of paracetamol

E. Coli metabolization of paracetamol


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For my science fair project, I did an experiment on how paracetamol and ibuprofen affected the growth of bacteria. By day 5, however, parts of the paracetamol blackened (image below). I am wondering if E. Coli can metabolize paracetamol and if so, what operon in the K-12 genome is responsible for this.

EDIT: I realized it may be metabolizing other ingredients in the medicine I used. I am including a list of the ingredients in the medicine I used.

Acetaminophen 160 mg in each 5 mL (duh)

Other ingredients (33.015873 mL):

  • Anhydrous citric acid
  • FD&C red no. 40
  • glycerin
  • high fructose corn syrup
  • microcrystalline cellulose and carboxymethylcellulose sodium
  • purified water
  • sodium benzoate
  • sorbitol solution
  • sucralose
  • xanthan gum

I thought it might be the fructose, but is the byproduct really black? Still confused…


Sorry that my answer is not a real answer, but this would be a good starting point for some more science.

Several of the questions you might ask and could answer (with some more experiments):

  • Does this happen in plates without E.coli?
  • You have a contamination on the right side, perhaps also on the black part. Could this be the culprit? Only half the plate is black, the rest is (still?) red.
  • You used a paracetamol tablet with red dye and a lot of sweeteners, does the same thing happen if you use one without the red dye? A dye would be a likely source, because it already contains an absorbing moiety. It only needs to be slightly modified to absorb different wavelengths.
  • What kind of agar are you using exactly? There are many kinds, often selecting for certain types of micro-organisms, with indicators for pH or other colored markers to detect and identify strains.

For your metabolic question you can take a look at https://ecocyc.org/ where you can find a database with all known metabolic pathways for E.coli K12. Unfortunately I couldn't find anything for paracetamol (or synonyms) but you could try to see if it produces a colored product for any of the additives.


Colorimetry: Principle and Instruments

Colorimetry is a widely used technique applied in biological system. It involves the measurement of a compound or a group of compounds present in a complex mixture. The property of colorimetric analyses is to determine the intensity or concentration of compounds in coloured solution.

This is done by passing light of specific wavelength of visible spectrum through the solution in a photoelectric colorimeter instrument and observe the galvanometric reading of reflection sensitizing the quantity of light absorbed.

Based on the nature of colour compounds, specific light filters are used. Three types of filters are available — blue, green and red — with corresponding light wavelength transmission rays from 470-490 nm, 500-530 nm and 620-680 nm, respectively.

There are two fundamental laws of absorption which are highly important in colorimetric estima­tion. These are Lambert’s law and Beer’s law. Lambert’s law states that when monochromatic light passes through a solution of constant concentration, the absorption by the solution is directly proportional to the length of the solution.

In contrary, Beer’s law states that when monochromatic light passes through a solution of constant length, the absorption by the solution is directly proportional to the concentration of the solution.

Thus both the laws can be expressed as:

[where I0 = Intensity of incident light (light entering a solution)

I = Intensity of transmitted light (light leaving a solution)

l = Length of absorbing solution

c = Concentration of coloured substance in solution

Both Beer-Lambert law are combined together for getting the expression transmittance (T).

(where I0 is the intensity of incident radiation and I is the intensity of transmitted radiation).

A 100% value of ‘T’ represent a totally transparent substance, with no radiation being aborted, whereas a zero value of ‘T’ represents a totally opaque substance that, in effect, represents complete absorption. For intermediate value we can define the absorbance (A) or extinction (E) that is given by the logarithm (to base 10) of the reciprocal of the transmittance:

Absorbance used to be called optical density (OD) but continued use of this term should be discour­aged. Also, as absorbance is a logarithm it is, by definition, unit-less and has a range of values from 0 (= 100% T) to cc (= 0% T).

Thus the variation of colour of the reaction mixture (or system) with change of substrate concentra­tion forms the basis of colorimetric analysis.

The formation of colour is due to the reaction between substances and reagents in appropriate proportion. The intensity of colour observed is then compared with that of reaction mixture which contains a known amount of substrate. The optical spectrophotometry is based on identical principles of colorimetry.

Instruments of Colorimetry:

The colorimeter instrument is very simple, consisting merely of a light source (lamp), filter, curette and photosensitive detector to collect the transmitted light. Another detector is required to measure the incident light or a single detector may be used to measure incident and transmitted light, alternately.

The latter design is both cheaper and analytically better, because it eliminates variations between detectors. The filter is used here to obtain an appropriate range of wavelengths within the bands which it is capable of selecting.

(B) Spectrophotometer:

It is a more sophisticated instrument. A photometer is a device for measuring ‘light’, and ‘spectro’ implies the whole range of continuous wavelengths that the light source is capable of producing. The detector in the photometer is generally a photo cell in which a sensitive surface receives photons and a current is generated that is proportional to the intensity of the light beam, reaching the surface.

In instru­ments for measuring ultraviolet/visible light, two lamps are usually required: one, a tungsten filament lamp which produces wavelengths in the visible region the second, a hydrogen or deuterium lamp, is suitable for the ultraviolet.

There are two kinds of optical arrangements: a single-beam or a double beam type. Here, first the blank and then the sample must be moved into the beam, adjustments made and readings taken.

The details of optical arrangement in Spectrophotometer is given:

The major advantage of the spectrophotometer, however, is the facility to scan the wavelength range over both ultraviolet and visible light and obtain absorption spectra.

A large number of inorganic and organic compounds were quantitatively estimated by the use of colorimetric or optical spectrophotometric techniques:


Evolving metabolism of 2,4-dinitrotoluene triggers SOS-independent diversification of host cells

The molecular mechanisms behind the mutagenic effect of reactive oxygen species (ROS) released by defective metabolization of xenobiotic 2,4-dinitrotoluene (DNT) by a still-evolving degradation pathway were studied. To this end, the genes required for biodegradation of DNT from Burkholderia cepacia R34 were implanted in Escherichia coli and the effect of catabolizing the nitroaromatic compound monitored with stress-related markers and reporters. Such a proxy of the naturally-occurring scenario faithfully recreated the known accumulation of ROS caused by faulty metabolism of DNT and the ensuing onset of an intense mutagenesis regime. While ROS triggered an oxidative stress response, neither homologous recombination was stimulated nor the recA promoter activity increased during DNT catabolism. Analysis of single-nucleotide changes occurring in rpoB during DNT degradation suggested a relaxation of DNA replication fidelity rather than direct damage to DNA. Mutants frequencies were determined in strains defective in either converting DNA damage into mutagenesis or mediating inhibition of mismatch repair through a general stress response. The results revealed that the mutagenic effect of ROS was largely SOS-independent and stemmed instead from stress-induced changes of rpoS functionality. Evolution of novel metabolic properties thus resembles the way sublethal antibiotic concentrations stimulate the appearance of novel resistance genes.


2 Photoactivatable Compounds and Optogenetic Proteins

Light-activation in biological systems can be either achieved through chemical modification with photosensitive groups and chemical effectors (chelators, isomers), or through genetically encoded photosensitive domains. The latter approach is referred to as optogenetic. This section discusses both approaches in general, and the properties of the individual components in particular as these lay the basis and set the limitations of engineering approaches for light-controllable systems. Chemical approaches such as photocaged molecules have been used decades before the first optogenetic methods were developed. However due to their flexibility and their unique dynamic properties, optogenetic regulators have quickly caught up as they are highly versatile for implementation in diverse cellular functions, and offer unique spatiotemporal control opportunities. The most important classes of light-sensitive protein modules for synthetic biology form the foundation for the discussion of potential engineering strategies for optogenetic regulators, and will be the main focus of this section.

2.1 Photoactivatable Compounds

Long before the term optogenetics was defined in 2006, optochemical approaches to measure and influence biological responses had been developed as early as the 1960s. This section gives an overview on photocageing groups, photosensitive chelators and cis-trans isomerization of azobenzenes. These serve as examples of some of the biologically relevant approaches, and this subsection is by no means intended to be comprehensive.

2.1.1 Photolabile Protecting Groups (PPGs)

Probably the most versatile of the three approaches is the use of PPGs (or photocaging groups) (Figure 2A). Although they have been used in different ways, the overall principle is that photocaged compounds contain a photolabile group, which renders a biomolecule inactive. Light induces the cleavage of the photolabile group which releases the biomolecule to its native function. [ 14 ] In 1962, Barltrop and Schofield [ 15 ] described the principle of PPGs. In 1977, Engels and Schlaeger described the synthesis and photolysis of an o-nitrobenzyl-caged cAMP and tested their activity using cAMP-dependent protein kinase. [ 16 ] One year later, Kaplan et al. [ 17 ] showed photolytic release of “caged ATP.” Since then, photocaging has been applied to numerous molecules from proteins to nucleic acids using a variety of photocleavable groups. For example, light can be used to remove protective groups in DNA synthesis, or fluorescently tagged photocleavable nucleotides in next generation sequencing approaches, such as sequencing by synthesis, or sequencing by ligation and microarray synthesis using photolithography and solid-phase synthesis. [ 14, 18-20 ]

The caged molecule varies depending on the cellular function that needs to be controlled with light. This molecule is rendered inactive through the caging group which contains a conjugated π system that can be cleaved-off upon photon stimulation. Two widely used photocaging groups are o-nitrobenzyl- and coumarin-based, which, depending on the substituents, can show absorption maxima from the UV to the green light spectrum. [ 21 ] Although many other caging groups exist, more than 80% of published photocaging approaches incorporate a nitrobenzyl group, which can be released in a well characterized photolysis mechanism. [ 14 ] Nitrobenzyl groups can be photocleaved using an excess of UVA light, even though the absorption maximum of most compounds lies in the UVB–UVC range, as UVA is considered far less damaging to cells than UVB–UVC. [ 14 ]

While numerous biofunctional chemicals have been caged (e.g., DOX, IPTG, arabinose, theophylline) with photosensitive groups (Figure 2A), the caging of proteins might show advantageous features: 1) the activity of cellular functions is precisely targeted through protein key players 2) only low concentrations compared to photocaged chemicals might avoid problems with photolysis byproducts. However, it needs to be considered that absorption of tryptophans could aid in the energy transfer and uncaging, and could also quench photolysis. In addition, pH and the local dielectric constant play an important role in the ground state absorption properties of a photolabile group, and one needs to take into account that for example the “apparent pKa” of a group can be different in the active site of an enzyme compared to other environments. Another difficulty can be the size, structure, and complexity of proteins, as photoreactive groups also need to be released after uncaging from an active site. [ 14 ] Overall, the advantages of photocaged proteins come with increased complexity in their synthesis.

Photosensitive groups can be introduced to proteins in vitro, for example through random modification with the oxycarbonylchloride of 1-(2-nitrophenyl)ethanol. The PPG primarily reacts with lysine residues, which was used to create light-activatable antibodies. [ 22 ] Another strategy targets cysteine residues as their nucleophilic groups allow selective modification by an electrophilic caging reagent. [ 14 ]

These and other strategies for in vitro synthesis of photoactivatable proteins can either be used in extracellular systems or need to be introduced into the cell using techniques such as microinjection, which can be prohibitive for many studies and applications. Therefore, in vivo synthesis of photoactivatable proteins can be enabling. An interesting approach is to expand the genetic code through the use of tRNA/aminoacyl-tRNA synthetase pairs, allowing one to include amino acids with photosensitive groups such as o-nitrobenzyl-caged tyrosine at amber sites. These caged amino acids can be located at functional sites of proteins such as the active site of an enzyme or the binding site of a protein. [ 23-25 ]

2.1.2 Photoresponsive Chelators

A different approach exploits chelators for light-control of cells by influencing the intracellular concentration of free metal ions, which fulfill numerous functions and are important cofactors for enzymes. This approach was very successful for buffers and optical indicators for Ca 2+ , which were synthesized based on BAPTA (l,2-bis(o-aminophenoxy)ethane-A,M-A/A'-tetraacetic acid). [ 26 ] This enabled experimenters to nondestructively measure intracellular Ca 2+ levels through shifts in the absorption spectrum of unbound to Ca 2+ -bound chelator (Figure 2B). [ 26 ] Ca 2+ ions have been of great interest for biological and biomedical studies, as it is an important component of cell signaling, and concentration changes are involved in diverse effects such as neuronal activity, cell motility, muscle contraction, apoptosis or transcription. [ 27 ] Although being able to measure intracellular Ca 2+ concentrations was effectively used for gaining biological insights, control of intracellular concentrations was highly desirable for perturbation studies. It was later realized that not only could concentrations be measured, but the affinity of the Ca 2+ chelators also changes with light. [ 28, 29 ] Chelator molecules, such as BAPTA, EDTA (ethylenediaminetetraacetic acid) or EGTA (ethylene glycol-bis(β-aminoethyl ether)-N,N,N′,N′-tetraacetic acid) form a “cavity” through the steric disposition of the carboxylate groups. [ 30 ] UV light is absorbed (maximum at 350 nm) by the caged compound and can lead to photolysis of the chelator to products that show a lower Ca 2+ affinity. Chelators for other divalent metal ions have also been developed. [ 31 ]

2.1.3 Cis–Trans Isomerization of Azobenzene

Cis–trans isomerization of azobenzene in response to light was described by Hartley in 1937. [ 32 ] This light-induced change in the structure of the chemical was later exploited to implement light control. It was shown that both forward and backward reactions are activated by light, and that the thermal reaction is slow. While the trans configuration, which is produced by 410–450 nm light, is planar, the benzene rings in the cis configuration are skewed at 53°, which can be triggered with 300–350 nm light. [ 29 ] Although the discovery of this phenomenon dates back more than 80 years, the underlying photochemistry and isomerization mechanism is still under investigation, with four proposed mechanisms: rotation, inversion, concerted inversion, and inversion-assisted rotation. [ 33 ] The isomerization can be repeated numerous times, as azobenzene shows high photostability, making it interesting for repeated photoswitching in dynamic control. The change in geometry can be used to modulate accessibility or activity of biomolecules. For example, it was shown that a 16 amino acid peptide tethered to an azobenzene can switch between a more helical structure in the trans, and a reduced helical content in the cis configuration (Figure 2C). [ 34 ] Tethered azobenzene was also used to controllably block and release the pore of a K + channel. Light with 380 nm wavelength, which creates the cis isomer, shortens the azobenzene and deblocks the pore, which allows K + ions to pass the channel, while 500 nm light creates the trans isomer which blocks the pore. [ 35 ]

2.2 Photoactivatable Proteins

As photocaged compounds show limited reversibility and spatial control once their protecting group is released, and as photoresponsive chelators and azobenzenes can be applied only in specific cases, new ways for light-control and the genetic implementations needed to achieve it, have been explored. Photoactivatable proteins or domains are essential components for optogenetic protein engineering. All photoactivatable domains used to-date in synthetic biology are derived from natural photoreceptors. Just like in other sensory signaling proteins, [ 36 ] modularity is common also in photosensors. In general, the input sensing domains or motifs are physically and functionally separable from the catalytic activity or the output domains. [ 36 ] This modularity feature of photosensory domains, which enables evolution through recombination, deletion, or insertion in the natural context, is also an enabling feature for bioengineering of novel light-inducible regulators. Indeed, such biological parts or modules with defined functionality are the fundamental basis for synthetic biology approaches. Through re-designing, combination with other modules, and engineering of the parts themselves, new functions can be created in organisms. [ 37 ] A prerequisite for the utilization of modules is their functional characterization, ideally with an understanding of the mechanism, the underlying structural basis, and the minimal requirements for the individual modules to work (e.g., availability of a specific cofactor or chromophore). As the understanding of photodomain properties is essential for development of functional optogenetic proteins, these properties are discussed in the following section.

The most widely used photoactivatable protein family in biology and medicine are light-sensitive transmembrane proteins. These proteins contain the chromophore retinal, or a variant, which in response to light isomerizes between an 11- or 13-cis and all-trans retinal. The chromophore isomerization translates to a structural change in the apoprotein. Depending on their origin, these so-called opsins are divided into microbial (Type I) and animal (Type II) opsins. The natural function of these proteins ranges from vision in animals, osmotic regulation in halobacteria, to photoperiodism in plants and animals. Although opsins have been used extensively in neurobiology and regenerative medicine for light-control of ion-fluxes and cell signaling, other non-opsin photoactivatable proteins lay the foundations for most protein engineering strategies in optogenetics. [ 38 ] Therefore, we present an overview of non-opsin photoactivatable proteins from nature, which have been adopted for synthetic biological approaches.

2.3 Chromophore and Photocycle

Photoactivatable proteins absorb light of specific wavelengths through an organic chromophore or cofactor which contains a conjugated π electron system. [ 39 ] Chromophores are molecules or chemical moieties that absorb light in the UV–vis spectrum. [ 40 ] The absorbed light leads to electron jumps from a lower to a higher energy molecular orbital, which in double-bonded molecules causes π–π * transitions. Conjugated π systems with conjugated electrons show a lower π–π * energy gap than single double-bonds and therefore absorb longer wavelengths and favor light absorption. [ 40 ] This chromophore excitation in turn leads to a structural change of the chromophore (e.g., cis–trans isomerization) and/or its interactions with the apoprotein, which leads to changes in the protein structure from a dark to a light activated state in a process called “photocycle.” This photocycle is closed once the activated photoactivatable protein reverts to the dark state. This dark state reversion is thermally driven, and its timescale can range from milliseconds to hours, depending on the photoreceptor.

2.4 Photosensor Classification

For the purpose of synthetic biology, we adopt an intuitive classification of photoactivatable proteins that is based on the protein's incorporated chromophore or the photoresponsive domain structure, which in part also determines the range of activation/inactivation wavelength peaks. Accordingly, four classes of photoactivatable proteins are described in this subsection: 1) light-activation via an intrinsic tryprophane 2) chromophores which are based on flavin, 3) cobalamin or 4) tetrapyrroles (Figure 3). Fluorescent proteins such as PhoCl, PYP or Dronpa, which were also used to implement light control are not discussed further. Our focus will be on the structural changes that light activation induces in the photoreceptor, as this is the basis for how they can be used in protein engineering approaches.

2.4.1 Intrinsic Tryptophan Regulated UVR8: UV Receptor

This photoreceptor class uses intrinsic tryptophanes for light absorption, with UVR8 as a prominent example. The photoregulator UVR8 was discovered in Arabidopsis thaliana as a mechanism to optimize growth and survival in the presence of UV-B. [ 41 ] UVR8 occurs as a homodimer in the dark state. [ 41 ] UV-B light is absorbed by tryptophane amino acid residues (W233, W285, and W337) which leads to monomerization of UVR8, and in turn allows for heterodimerization with UVR8-binding partner COP1. [ 41 ] W233 and W285, which act as UV-B chromophore, show cation–π interactions with R286 and R338 and stabilize the protein structure. Excitation of the tryptophan indole rings through UV-B light disrupts these interactions, which leads to the release of arginine-mediated intermolecular hydrogen bonds between the homodimers and UVR8 monomerization. [ 42 ] The photocycle is closed as the indole rings dissipate energy and return to their ground state over time, leading to homodimerization of UVR8. [ 42 ]

2.4.2 Flavin-Based Cryptochromes, BLUF, and LOV Domains: Blue Light Receptors

Cryptochromes, LOV, and BLUF domains contain a flavin chromophore, either flavin mononucleotide (FMN) or flavin adenine dinucleotide (FAD), which can be covalently or noncovalently bound to the apoprotein. [ 43 ] FMN and FAD are present in most organisms. Flavin shows maximum absorption in the blue light range which causes a photochemical reduction of the oxidized form to a semiquinone or the fully reduced form. [ 44, 45 ] The isoalloxazine ring system allows for one- or two electron transfer processes. [ 43 ] Also multi-photon excitation with near-infrared light was shown, allowing for 3D activation of flavin-based systems and deep tissue penetration. [ 46-48 ]

The evolutionarily highly conserved cryptochromes CRY1 and CRY2 belong to the family of flavoproteins that exist in all kingdoms of life, where they are involved in developmental and circadian responses. [ 49 ] Cryptochromes evolved from photolyases and contain an N-terminal photolyase homology (PHR) domain, which binds FAD as chromophore. [ 49 ] In a proposed light-activation mechanism in plant cryptochrome, oxidized FAD in the dark state is reduced to a neutral semiquinone through light, which induces a negative charge in the vicinity of the flavin. This might lead to the release of ATP from its binding pocket, and the subsequent C-terminus unfolding of the protein. Through the conformational change and release of the C-terminus, amino acid residues are accessible for phosphorylation, which then allows for binding of other proteins. [ 50 ] A widely used cryptochrome is CRY2 from Arabidopsis thaliana (AtCRY2). CRY2 is monomeric in the dark state, and oligomerizes upon blue light activation. [ 51 ] In the photoexcited state, it can also form a heterodimer with cryptochrome-interacting basic-helix-loop-helix protein (CIB1). [ 52 ] The half-life of this interaction is in the minute-scale, and it can be tuned through modulating mutations in the PHR domain. [ 53 ]

The second class of sensors that absorb blue light using FAD (BLUF domain) was discovered in Rhodobacter sphaeroides [ 54, 55 ] and Euglena gracilis, [ 56 ] where they aid in adaptation of photosystem synthesis depending on oxygen and light conditions. [ 57 ] Although protein structures of BLUF domain containing proteins have been solved, the mechanism of photo-activation is still under debate. [ 57, 58 ] Through blue light-illumination, an electron and then a proton is transferred from a conserved tyrosine to the flavin, which leads to formation of flavin and tyrosine radicals. [ 48, 59 ] The bi-radical then might induce a hydrogen bond rearrangement in the flavin binding pocket. [ 57 ] PixD is an example of a pentameric BLUF domain that, together with PixE, form large molecular weight aggregates of two pentameric PixD and 5 PixE subunits. [ 60 ] Through illumination, this complex can be destabilized, resulting in monomeric PixE, and two pentameric PixD. [ 60 ]

Another widely used class of blue light-sensitive proteins are flavin mononucleotide (FMN)-binding light oxygen and voltage (LOV) domains. In contrast to cryptochromes, where light causes electrostatic changes in the apoprotein and subsequent conformational changes, the FMN C(4a) in LOV domains usually forms a covalent adduct with an adjacent conserved cysteine, which in turn also causes conformational changes in the Per-ARNT-Sim (period clock protein, aromatic hydrocarbon receptor nuclear translocator, and single minded short PAS) core. [ 61-63 ] Apart from FMN, FAD functions also as a chromophore in LOV domains, as for example in the photoregulator Vivid. [ 64 ] PAS domains are found in all kingdoms of life and commonly act as molecular sensors and transducers. [ 65, 66 ] The generated structural changes through cysteinyl-(C4a) formation propagate to N- or C-terminally attached effector domains via amphipathic α-helical and coiled-coil linkers. This conformational change can initiate diverse mechanisms such as dimerization of LOV domains in Vivid, [ 62, 64, 67 ] the unfolding and displacement of a Jα helix in the case of AsLOV2, [ 66 ] or the rearrangement and release of a helix-turn-helix (HTH) domain for EL222 [ 68 ] from their respective LOV core. Based on these characteristics, LOV1 domains usually involve a PAS core in some cases with an additional N-terminal cap (NCap) through which the proteins associate. [ 69-71 ] LOV2 domains, on the other hand, contain a C-terminal Jα helix which displaces after light-stimulation. [ 72, 73 ]

2.4.3 Cobalamin-Based Binding Domains: Green Light Receptors

Cobalamin-binding domains (CBDs) are green light photoreceptors that utilize cobalamin as chromophore for photosensing. CBDs were found to play a photoprotecting role in diverse bacteria [ 74-76 ] and were identified through their role in light-dependent carotenoid synthesis which quenches reactive oxygen species (ROS). [ 77-79 ] One of the CBDs that were used so far in synthetic biology is CarH. The CarH photoreceptor dimer binds 5′deoxyadenosylcobalamin (AdoCbl) as its chromophore to form a tetramer. AdoCbl and methylcobalamin (MeCbl) are the two major biological forms of Vitamin B12 which are produced by microorganisms. [ 80 ] Mammalian cells are capable of AdoCbl import and its conversion from Vitamin B12. [ 81, 82 ] AdoCbl and MeCbl differ in the 5′deoxyadenosyl and methyl group that is covalently bound to cobalt, which in both cases has low bond dissociation energies. The low dissociation energies allow for the cleavage of the respective 5′deoxyadenosyl or methyl groups with wavelengths ranging from near-UV light up to wavelengths of 530 nm in the green light spectrum. [ 80 ] In the case of CarH, the proteins form head-to-tail tetramers as a dimer-of-dimers in the presence of AdoCbl which involves numerous hydrogen bonds of the apoprotein with cobalamin and hydrogen bonds and ionic interactions with the 5′deoxyadenosyl group. [ 83 ] Light exposure triggers dissociation of the 5′deoxyadenosyl group, which leads to reorientation of the four-helix bundle of the protein disrupting the head-to-tail interface, resulting in CarH monomerization. [ 83 ] Since the cobalamin forms a covalent adduct with CarH through bis-His ligation, this reaction is irreversible, and photolyzed cobalamin cannot be exchanged with photosensitive AdoCbl. [ 83 ] This limits applicability of CarH for fast dynamic optogenetic control.

2.4.4 Tetrapyrrole-Based Phytochromes: From UV to Far-Red Receptors

Apart from flavin-based photoreceptors, phytochromes are the second widely used class of light-inducible domains which were used in the first optogenetic regulators in eukaryotic [ 2 ] and bacterial [ 84 ] cells. Phytochromes (Phy) were discovered through their role in promoting development in plants such as germination and flowering in response to red-light. [ 85-87 ] However, phytochromes are not just present in plants, but also in bacteria [ 88, 89 ] (BphP) and fungi [ 90 ] (Fph). Phytochromes are classified depending on their activation light wavelength to type I, which is activated by far-red light (730 nm absorption peak), and type II, which is activated by red light (660 nm absorption peak). The two types therefore differ in their thermal ground states and can reversibly switch between a red-absorbing (Pr) and a far-red-absorbing (Pfr) state. However, algal phytochromes [ 91 ] and cyanobacteriochromes [ 92 ] were described and shown to cover the full visible spectrum and even reach into the UV range. For example, phytochromes with blue(Pb)–green(Pg) [ 93 ] and green(Pg)–red(Pr) [ 94 ] -activating wavelengths were discovered. Even though they absorb a wide range of wavelengths, all phytochromes use a tetrapyrrole chromophore, either in the reduced form as phycocyanobilin (i.e., plant photochromes and cyanobacteriochromes), or in the oxidized form as biliverdin as used by bacterial and fungal phytochromes. [ 95 ] Biliverdin (BV), the chromophore for BphP and Fph, is synthesized from heme in one step through heme oxygenase HO1. In cyanobacteria and green algae, BV is further reduced to phycocyanobilin (PCB) by a ferredoxin-dependent bilin reductase PcyA. In plants however, an enzyme of the same family called HY2 reduces BV to phytochromobilin (PΦB). [ 95-97 ] The different bilins are bound in a GAF domain, which is highly conserved. Typical phytochromes show a PAS-GAF-PHY domain photosensory module structure, although variations in this structure exist. [ 95 ] In a suggested photoactivation mechanism, light induces a Z–E isomerization in the C15-C16 double bond of the tetrapyrrole, [ 98 ] inducing a rotation on the D-ring of the molecule, which in turn generates a rearrangement of the hydrogen bonds of the GAF domain that propagates to the PHY domain. [ 95, 99 ] These structural changes can have diverse effects in different phytochromes and, for example, allow for heterodimerization of light-induced PhyB with phytochrome-interacting factor (PIF3), or homodimerization in the case of cyanobacterial phytochrome Cph1. [ 88 ] Furthermore, phytochromes do not just rely on dark state reversion over time through energy dissipation, but they can be induced specifically with deactivating light at a wavelength different from the induction wavelength (e.g., far-red light for PhyB [ 100 ] or Cph1 [ 88 ] ). This gives phytochromes a superior temporal resolution.


Abstract

N-ε-Carboxymethyllysine (CML) is formed during glycation reactions (synonym, Maillard reaction). CML is degraded by the human colonic microbiota, but nothing is known about the formation of particular metabolites. In the present study, six probiotic E. coli strains were incubated with CML in the presence or absence of oxygen in either minimal or nutrient-rich medium. CML was degraded by all strains only in the presence of oxygen. HPLC-MS/MS was applied for identification of metabolites of CML. For the first time, three bacterial metabolites of CML have been identified, namely N-carboxymethylcadaverine (CM-CAD), N-carboxymethylaminopentanoic acid (CM-APA), and the N-carboxymethyl-Δ 1 -piperideinium ion. During 48 h of incubation of CML with five different E. coli strains in minimal medium in the presence of oxygen, 37–66% of CML was degraded, while CM-CAD (1.5–8.4% of the initial CML dose) and CM-APA (0.04–0.11% of the initial CML dose) were formed linearly. Formation of the metabolites is enhanced when dipeptide-bound CML is applied, indicating that transport phenomena may play an important role in the “handling” of the compound by microorganisms.


Lab 12: Isolation and Identification of Enterobacteriaceae and Pseudomonas, Part 1

Labs 12 and 13 deal with opportunistic and pathogenic fermentative Gram-negative bacilli that are members of the bacterial family Enterobactereaceae, as well as nonfermentative Gram-negative bacilli such as Pseudomonas and Acinetobacter.

A. ENTEROBACTERIACEAE: THE FERMENTATIVE, GRAM-NEGATIVE, ENTERIC BACILLI

Bacteria belonging to the family Enterobacteriaceae are the most commonly encountered organisms isolated from clinical specimens. The Enterobacteriaceae is a large diverse family of bacteria belonging to the order Enterobacteriales in the class Gammaproteobacter of the phylum Proteobacter. Medically important members of this family are commonly referred to as fermentative, Gram-negative, enteric bacilli, because they are Gram-negative rods that can ferment sugars. Many are normal flora of the intestinal tract of humans and animals while others infect the intestinal tract. Members of this family have the following characteristics in common:

1. They are Gram-negative rods (see Fig. 1)
2. If motile, they possess a peritrichous arrangement of flagella (see Fig. 2)
3. They are facultative anaerobes
4. With few exception, they are oxidase negative
5. All species ferment the sugar glucose but otherwise vary widely in their biochemical characteristics
6. Most reduce nitrates to nitrites.

For further information on the Gram-negative cell wall, see the following Learning Object in your Lecture Guide:

At least forty-four genera and over 130 species of Enterobacteriaceae have been recognized. Some of the more common clinically important genera of the family Enterobacteriaceae include:

Salmonella Citrobacter Morganella
Shigella Enterobacter Yersinia
Proteus Serratia Edwardsiella
Escherichia Klebsiella Providencia

Several genera of Enterobacteriaceae are associated with gastroenteritis and food-borne disease. These include:

  • Salmonella,
  • Shigella,
  • certain strains of Escherichia coli, and
  • certain species of Yersinia.

All intestinal tract infections are transmitted by the fecal-oral route.

There are two species of Salmonella, Salmonella enterica and Salmonella bongori. Any infection caused by Salmonella is called a salmonellosis. Non-typhoidal Salmonella accounts for an estimated 520 cases per 100,000 population (approximately 1,600,000 cases) per year in the U.S. and at least 500 die. Since many different animals carry Salmonella in their intestinal tract, people usually become infected from ingesting improperly refrigerated, uncooked or undercooked poultry, eggs, meat, dairy products, vegetables, or fruit contaminated with animal feces.

E nteritis is the most common form of salmonellosis. Symptoms generally appear 6-48 hours after ingestion of the bacteria and include vomiting, nausea, non-bloody diarrhea, fever, abdominal cramps, myalgias, and headache. Symptoms generally last from 2 days to 1 week followed by spontaneous recovery. All species of Salmonella can cause bacteremia but S. enterica serotype Typhi, isolated only from humans, frequently disseminates into the blood causing a severe form of salmonellosis called typhoid fever. About 400 cases of typhoid fever occur each year in the U.S. but approximately 75% of these are acquired while traveling internationally.

Salmonella serotyping is a subtyping method of identification based on the identification of distinct cell wall, flagellar, and capsular antigens with known antiserum, as will be discussed in Lab 17. Salmonella serotypes Enteritidis and Typhimurium are the two most common serotypes in the United States, accounting for approximately 35 to 40% of all infections confirmed by laboratory culture. As mentioned above, S. enterica serotype Typhi is responsible for typhoid fever.

Any Shigella infection is called a shigellosis. Unlike Salmonella, which can infect many different animals, Shigella only infects humans and other higher primates. There are approximately 14,000 laboratory cases of shigellosis a year reported in the US with an estimated 450,000 total cases and 70 deaths.

Shigellosis frequently starts with a watery diarrhea, fever, and abdominal cramps but may progress to dysentery with scant stools containing blood, pus, and mucus. The incubation period is 1-3 days. Initial profuse watery diarrhea typically appears first as a result of enterotoxin. Within 1-2 days this progresses to abdominal cramps, with or without bloody stool. Classic shigellosis presents itself as lower abdominal cramps and stool abundant with blood and pus develops as the Shigella invade the mucosa of the colon.

Escherichia coli is one of the dominant normal flora in the intestinal tract of humans and animals. Some strains, however, can cause infections of the intestines while others are capable of causing infections outside the intestines. Extraintestinal pathogenic E. coli cause such opportunistic infections as urinary tract infections, wound infections, and septicemia and will be discussed in greater detail below. Intestinal or diarrheagenic E. coli cause infections of the intestinal tract. Diarrheagenic E. coli include:

  • Enterotoxigenc E. coli (ETEC) produce enterotoxins that cause the loss of sodium ions and water from the small intestines resulting in a watery diarrhea. It is an important cause of diarrhea in impoverished countries. Over half of all travelers' diarrhea is due to ETEC almost 80,000 cases a year in the U.S.
  • Enteropathogenic E. coli (EPEC) causes an endemic diarrhea in in impoverished countries, especially in infants younger than 6 months of age. The bacterium disrupts the normal microvilli on the epithelial cells of the small intestines resulting in maladsorbtion and diarrhea. They do not produce enterotoxin or shiga toxin and are not invasive. It is rare in industrialized countries.
  • Enteroaggregative E. coli (EAEC) is a cause of endemic diarrhea in children in impoverished countries and industrialized countries. It is also responsible for a persistant diarrhea in people infected with HIV. It probably causes diarrhea by adhering to mucosal epithelial cells of the small intestines and interfering with their function.
  • Enteroinvasive E. coli (EIEC) invade and kill epithelial cells of the colon usually causing a watery diarrhea but sometimes progressing to a dysentery-type syndrome with blood in the stool. It occurs mostly in impoverished countries and is rare in industrialized countries.
  • Enterohemorrhagic E. coli (EHEC), such as E. coli 0157:H7, produce a shiga toxin that kills epithelial cells of the colon causing hemorrhagic colitis, a bloody diarrhea. In rare cases, the shiga toxin enters the blood and is carried to the kidneys where, usually in children, it damages vascular cells and causes hemolytic uremic syndrome. E. coli 0157:H7 is thought to cause more than 20,000 infections and up to 250 deaths per year in the U.S.

Several species of Yersinia, such as Y. enterocolitica and Y. pseudotuberculosis are also causes of diarrheal disease.

Many other genera of the family Enterobacteriaceae are normal microbiota of the intestinal tract and are considered opportunistic pathogens. The most common genera of Enterobacteriaceae causing opportunistic infections in humans are:

  • Escherichia coli,
  • Proteus,
  • Enterobacter,
  • Klebsiella,
  • Citrobacter, and
  • Serratia.

They act as opportunistic pathogens when they are introduced into body locations where they are not normally found, especially if the host is debilitated or immunosuppressed. They all cause the same types of opportunistic infections, namely:

  • urinary tract infections,
  • wound infections,
  • pneumonia, and
  • septicemia.

These normal flora Gram-negative bacilli, along with Gram-positive bacteria such as Enterococcus species (see Lab 14) and Staphylococcus species (see Lab 15), are among the most common causes of healthcare-associated infections (formerly called nosocomial infections).

According to the Centers for Disease Control and Prevention (CDC) Healthcare-associated infection's website, "In American hospitals alone, healthcare-associated infections account for an estimated 1.7 million infections and 99,000 associated deaths each year. Of these infections:

  • 32 percent of all healthcare-associated infection are urinary tract infections (UTIs)
  • 22 percent are surgical site infections
  • 15 percent are pneumonia (lung infections)
  • 14 percent are bloodstream infections"

Most patients who have healthcare-associated infections are predisposed to infection because of invasive supportive measures such as urinary catheters, intravascular lines, and endotracheal intubation.

By far, the most common Gram-negative bacterium causing nosocomial infections is Escherichia coli. E. coli causes between 70 and 90% of both upper and lower urinary tract infections (UTIs). It is also a frequent cause of abdominal wound infections and septicemia. Depending on the facility, E. coli is responsible for between 12% and 50% of all healthcare-associated infections.

However, according to a 2008 study, Enterobacteriaceae other than E. coli were responsible for 7 of the 10 most common Gram-negative organisms isolated from urinary tract, respiratory tract, and bloodstream infections from intensive care unit patients between 2002 and 2008 in the United States. These include Klebsiella pneumoniae (15%), Enterobacter cloacae (9%), Serratia marcescens (6%), Enterobacter aerogenes (4%), Proteus mirabilis (4%), Klebsiella oxytoca (3%), and Citrobacter freundii (2%). Furthermore, the National Healthcare Safety Network reported K. pneumoniae (6%) , Enterobacter spp. (5%), and K. oxytoca (2%) among the top 10 most frequently isolated health care-associated infections between the years between 2006 and 2007.

1. Urinary Tract Infections

The most common infection caused by opportunistic Enterobacteriaceae is a urinary tract infection (UTI). UTIs account for more than 8, 000,000 physician office visits per year in the U.S and as many as 100,000 hospitalizations. Among the nonhospitalized and nondebilitated population, UTIs are more common in females because of their shorter urethra and the closer proximity between their anus and the urethral opening. (Over 20 percent of women have recurrent UTIs.) However, anyone can become susceptible to urinary infections in the presence of predisposing factors that cause functional and structural abnormalities of the urinary tract. These abnormalities increase the volume of residual urine and interfere with the normal clearance of bacteria by urination. Such factors include prostate enlargement, sagging uterus, expansion of the uterus during pregnancy, paraplegia, spina bifida, scar tissue formation, and catheterization. Between 35 and 40 percent of all nosocomial infections, about 900,000 per year in the U.S., are UTIs and are usually associated with catheterization.

E. coli and Staphylococcus saprophyticus (a Gram-positive staphylococcus that will be discussed in Lab 15) cause around 90 percent of all uncomplicated UTIs. Most of the remaining uncomplicated UTIs are caused by other Gram-negative enterics such as Proteus mirabilis and Klebsiella pneumoniae or by Enterococcus faecalis (a Gram-positive streptococcus that will be discussed in Lab 14). E. coli is responsible for more than 50 percent of healthcare-associated UTIs. Other causes of hospital-acquired UTIs include other species of Enterobacteriaceae (such as Proteus, Enterobacter, and Klebsiella), Pseudomonas aeruginosa (discussed below), Enterococcus species (discussed in lab 14) , Staphylococcus saprophyticus (discussed in Lab 15), and the yeast Candida (discussed in lab 9).

The traditional laboratory culture standard for a UTI has been the presence ofmore than 100,000 CFUs (colony-forming units see Lab 4) per milliliter (ml) of midstream urine, or any CFUs from a catheter-obtained urine sample. More recently, this has been modified and counts of as few as 1000 colonies of a single type per ml or as little as 100 coliforms per ml are now considered as indicating a UTI.

2. Wound Infections

Wound infections are due to fecal contamination of external wounds or a result of wounds that cause trauma to the intestinal tract, such as surgical wounds, gunshot wounds, and knife wounds. In the latter case, fecal bacteria get out of the intestinal tract and into the peritoneal cavity causing peritonitis and formation of abcesses on the organs found in the peritoneal cavity.

3. Pneumonia

Although they sometimes cause pneumonia, the Enterobacteriaceae account for less than 5% of the bacterial pneumonias requiring hospitalization.

4. Bloodstream Infections

Gram-negative septicemia is a result of these opportunistic Gram-negative bacteria getting into the blood. They are usually introduced into the blood from some other infection site, such as an infected kidney, wound, or lung. Looking at patients that develop septic shock:

  • Lower respiratory tract infections are the source in about 25% of patients.
  • Urinary tract infections are the source in about 25% of patients.
  • Soft tissue infections are the source in about 15% of patients.
  • Gastrointestinal infections are the source in about 15% of patients.
  • Reproductive tract infections are the source in about 10% of patients.
  • Foreign bodies (intravascular lines, implanted surgical devices, etc.) are the source in about 5% of patients.

There are approximately 750,000 cases of septicemia per year in the U.S. and 200,000 cases of septic shock. Septic shock results in approximately 100,000 deaths per year in the U.S. Approximately 45 percent of the cases of septicemia are due to Gram-negative bacteria. Klebsiella, Proteus, Enterobacter, Serratia, and E. coli, are all common Enterobacteriaceae causing septicemia. (Another 45 percent are a result of Gram-positive bacteria (see Labs 14 and 15) and 10 percent are due to fungi, mainly the yeast Candida (see Lab 9).

In the outer membrane of the Gram-negative cell wall, the lipid A moiety of the lipopolysaccharide (LPS) functions as an endotoxin (see Fig 4 ). Endotoxin indirectly harms the body when massive amounts are released during severe Gram-negative infections. This, in turn, causes an excessive cytokine response.

1. TheLPS released from the outer membrane of the Gram-negative cell wall first binds to a LPS-binding protein circulating in the blood and this complex, in turn, binds to a receptor molecule (CD 14 ) found on the surface of body defense cells called macrophages (see Fig. 5) located in most tissues and organs of the body.

2. This is thought to promote the ability of the toll-like receptor TLR-4 to respond to the LPS, triggering the macrophages to release various defense regulatory chemicals called cytokines, including tumor necrosis factor-alpha (TNF-alpha), interleukin-1 (IL-1), interleukin-6 (IL-6), and interleukin-8 (IL-8), and platelet-activating factor (PAF). The cytokines then bind to cytokine receptors on target cells and initiate inflammation and activate both the complement pathways and the coagulation pathway (see Fig. 5).

3. The complex of LPS and LPS binding protein can also attach to molecules called CD14 on the surfaces of phagocytic white blood cells called neutrophils causing them to release proteases and toxic oxygen radicals for extracellular killing. Chemokines such as interleukin-8 (IL-8) also stimulate extracellular killing. In addition, LPS and cytokines stimulate the synthesis of a vasodilator called nitric oxide.

D uring minor local infections with few bacteria present, low levels of LPS are released leading to moderate cytokine production by the monocytes and macrophages and in general, promoting body defense by stimulating inflammation and moderate fever, breaking down energy reserves to supply energy for defense, activating the complement pathway and the coagulation pathway, and generally stimulating immune responses (see Fig. 5). Also as a result of these cytokines, circulating phagocytic white blood cells such as neutrophils and monocytes stick to the walls of capillaries, squeeze out and enter the tissue, a process termed diapedesis. The phagocytic white blood cells such as neutrophils then kill the invading microbes with their proteases and toxic oxygen radicals.

However, during severe systemic infections with large numbers of bacteria present, high levels of LPS are releasedresulting inexcessive cytokine productionby the monocytes and macrophages and this canharm the body (see Fig. 6). In addition, neutrophils start releasing their proteases and toxic oxygen radicals that kill not only the bacteria, but the surrounding tissue as well. Harmful effects include high fever, hypotension, tissue destruction, wasting, acute respiratory distress syndrome (ARDS), disseminated intravascular coagulation (DIC), and damage to the vascular endothelium resulting in shock, multiple system organ failure (MOSF), and often death.

This excessive inflammatory response is referred to as Systemic Inflammatory Response Syndrome or SIRS. Death is a result of what is called the shock cascade. The sequence of events is as follows:

  • Neutrophil-induced damage to the capillaries, as well as prolonged vasodilation, results in blood and plasma leaving the bloodstream and entering the surrounding tissue. This can lead to a decreased volume of circulating blood (hypovolemia).
  • Prolonged vasodilation also leads to decreased vascular resistance within blood vessels while high levels of TNF inhibit vascular smooth muscle tone and myocardial contractility. This results in a marked hypotension.
  • Activation of the blood coagulation pathway can cause clots called microthrombi to form within the blood vessels throughout the body causing disseminated intravascular coagulation (DIC).
  • Increased capillary permeability as a result of vasodilation in the lungs, as well as neutrophil-induced injury to capillaries in the alveoli, lead to acute inflammation, pulmonary edema, and loss of gas exchange in the lungs (acute respiratory distress syndrome or ARDS). As a result, the blood does not become oxygenated.
  • Hypotension, hypovolemia, ARDS, and DIC result in marked hypoperfusion.
  • Hypoperfusion in the liver can result in a drop in blood glucose level from liver dysfunction.
  • Hypoperfusion leads to acidosis and the wrong pH for enzymes involved in cellular metabolism resulting in cell death.
  • Hypoperfusion also can lead to cardiac failure .

Collectively, this can result in :

  • end-organ ischemia: a restriction in blood supply that results in damage or dysfunction of tissues or organs,
  • multiple system organ failure (MSOF),
  • death.

Both pili and surface proteins in the Gram-negative cell wall function as adhesins, allowing the bacterium to adhere intimately to host cells and other surfaces in order to colonize and resist flushing. Some Gram-negative bacteria also produce invasins, allowing some bacteria to invade host cells. Motility, capsules, biofilm formation, and exotoxins also play a role in the virulence of some Enterobacteriaceae.

For further information on virulence factors associated with various Enterobacteriaceae, see the following Learning Objects in your Lecture Guide:

Many of the Enterobacteriaceae also possess R (resistance) plasmids. These plasmids are small pieces of circular non-chromosomal DNA that may code for multiple antibiotic resistance In addition, the plasmid may code for a sex pilus, enabling the bacterium to pass R plasmids to other bacteria by conjugation. Between 50 and 60 percent of the bacteria causing healthcare-associated infections are antibiotic resistant.

For further information on bacterial resistance to antibiotics, see the following Learning Object in your Lecture E-Text:

The identification of lactose-fermenting Gram-negative rods belonging to the bacterial family Enterobacteriaceae (bacteria commonly referred to as coliforms) in water is often used to determine if water has been fecally contaminated and, therefore, may contain disease-causing pathogens transmitted by the fecal-oral route. The procedure for this is given in Appendix E.

B. PSEUDOMONAS AND OTHER NON-FERMENTATIVE GRAM-NEGATIVE BACILLI

Non-fermentative Gram-negative bacilli refer to Gram-negative rods or coccobacilli that cannot ferment sugars. The non-fermentative Gram-negative bacilli are often normal inhabitants of soil and water. They may cause human infections when they colonize immunosuppressed individuals or gain access to the body through trauma. However, less than one-fifth of the Gram-negative bacilli isolated from clinical specimens are non-fermentative bacilli. By far, the most common Gram-negative, non-fermentative rod that causes human infections is Pseudomonas aeruginosa. Pseudomonas belongs to the family Pseudomonadaceae in the order Pseudomonadales in the class Gammaproteobacter of the phylum Proteobacter.

Pseudomonas aeruginosa is also an opportunistic pathogen. It is a common cause of nosocomial infections and can be found growing in a large variety of environmental locations. In the hospital environment, for example, it has been isolated from drains, sinks, faucets, water from cut flowers, cleaning solutions, medicines, and even disinfectant soap solutions. It is especially dangerous to the debilitated or immunocompromised patient.

Like the opportunistic Enterobacteriaceae, Pseudomonas is a Gram-negative rod, it is frequently found in small amounts in the feces, and it causes similar opportunistic infections: urinary tract infections, wound infections, pneumonia, and septicemia. P aeruginosa is the fourth most commonly isolated nosocomial pathogen, accounting for 10% of all hospital-acquired infections. P. aeruginosa is responsible for 12 percent of hospital-acquired urinary tract infections, 16 percent of nosocomial pneumonia cases, and 10 percent of the cases of septicemia. In addition, P. aeruginosa is a significant cause of burn infections with a 60 percent mortality rate. It also colonizes and chronically infects the lungs of people with cystic fibrosis. Like other opportunistic Gram-negative bacilli, Pseudomonas aeruginosa also releases endotoxin and frequently possesses R-plasmids. A number of other species of Pseudomonas have also been found to cause human infections.

For further information on virulence factors associated with Pseudomonas, see the following Learning Objects in your Lecture Guide:

Other non-fermentative Gram-negative bacilli that are sometimes opportunistic pathogens in humans include Acinetobacter, Aeromonas, Alcaligenes, Eikenella, Flavobacterium, and Moraxella.

Acinetobacter has become a frequent cause of nosocomial wound infections, pneumonia, and septicemia. The bacterium has become well known as a cause of infections among veterans of the wars in Iraq and Afghanistan and is becoming a growing cause of nosocomial infections in the U.S. Acinetobacter is thought to have been contracted in field hospitals in Iraq and Afghanistan and subsequently carried to veteran's hospitals in the U.S. Because most species are multiple antibiotic resistant, it is often difficult to treat. Acinetobacter is commonly found in soil and water, as well as on the skin of healthy people, especially healthcare personnel. Although there are numerous species of Acinetobacter that can cause human disease, Acinetobacter baumannii accounts for about 80% of reported infections.

Medscape articles on infections associated with organisms mentioned in this lab exercise. Registration to access this website is free.

  • Salmonellosis
  • Typhoid fever
  • Shigellosis
  • Escherichia coli
  • Proteusspecies
  • Klebsiellaspecies
  • Enterobacterspecies
  • Serratiaspecies
  • Yersinia enterocolitica
  • Yersinia pseudotuberculosis
  • Acinetobacter baumannii
  • Pseudomonas aeruginosa
  • Urinary tract infections
  • Wound infections
  • Community-acquired pneumonia
  • Sepsis

SCENERIOS FOR TODAY'S LAB

Students will be assigned either Case Study 1A or 1B to do today. All students will do Case Study 2 as part of the results next time.

Case Study #1A

A 66 year old female with a history of recurring urinary tract infections and multiple antibiotic therapies presents with frequency and urgency of urination, dysuria, suprapubic discomfort, lower back pain, and a temperature of 99.2°F. A complete blood count (CBC) shows leukocytosis with a left shift. A urine dipstick shows a positive leukocyte esterase test, a positive nitrite test, 30mg of protein per deciliter, and red blood cells in the urine.

Assume that your unknown is a urine culture from this person.

Case Study #1B

A 72 year old female who is diabetic and a smoker was admitted to the hospital with a leg wound that is not healing. She appears confused and anxious, has a temperature of 102 °F , a heart rate of 101 beats per minute, a respiration rate of 29 breaths per minute, a blood pressure of 94/32 mm Hg, a urine output of only 110 cc for the last 8 hours, and a total white blood cell count of of 2300/µL. A blood culture is taken.

Assume that your unknown is a blood culture from this person.

CAUTION: TREAT EACH UNKNOWN AS A PATHOGEN!. Inform your instructor of any spills or accidents. WASH AND SANITIZE YOUR HANDS WELL before leaving the lab.

Taxo N® disk, alcohol, dropper bottle of distilled water, swab, and either a plate of MacConkey agar or a plate of Cetrimide agar, and an EnteroPluri-Test

PROCEDURE (to be done in groups of 3 )

[Keep in mind that organisms other than the Enterobacteriaceae and Pseudomonas can cause these infections, so in a real clinical situation other lab tests and cultures for bacteria other than those upon which this lab is based would also be done.]

1 . Perform a Gram stain on your unknown. Remember that the concentration of bacteria on slides prepared from taking bacteria off a petri plate tend to be much greater than those prepared by taking bacteria out of a broth culture, so be careful not to under decolorize. Continue decolorizing until the purple just stops flowing off of the bacterial smear, then wash with water.

Record the results of your Gram stain in the Gram stain section of Lab 13.

2. If you have a Gram-negative bacillus, determine if it is a fermentative Gram-negative bacillus like most Enterobacteriaceae or a non-fermentative Gram-negative bacillus such as Pseudomonas by performing an oxidase test as follows:

a. Using alcohol-flamed forceps, remove a Taxo-N® disc and moisten it with a drop of sterile distilled water.

b. Place the moistened disc on the colonies of the culture of your unknown.

c. Using a sterile swab, scrape off some of the colonies and spread them on the Taxo-N® disc.

In the immediate test, oxidase-positive reactions will turn a rose color within 30 seconds (see Fig. 10). Oxidase-negative will not turn a rose color (see Fig. 9). This reaction only lasts a couple of minutes. In the delayed test, oxidase-positive colonies within 10 mm of the Taxo-N® disc will turn black within 20 minutes and will remain black (see Fig 11). If the bacterium is oxidase-negative, the growth around the disc will not turn black (see Fig. 12).

Record your oxidase test results in the Oxidase test section of Lab 13.

3. If your unknown is oxidase-negative, indicating a fermentative Gram-negative bacillus, do the following inoculations:

a. Streak your unknown for isolation on a plate of MacConkey agar, a selective medium used for the isolation of non-fastidious Gram-negative rods and particularly members of the family Enterobacteriaceae, using one of the two streaking patterns illustrated in Fig. 4 and Fig. 5. Incubate upside down and stacked in the petri plate holder on the shelf of the 37°C incubator corresponding to your lab section.

b. Inoculate an EnteroPluri-Test as follows:

1. Remove both caps of the EnteroPluri-Test and with the straight end of the inoculating wire, pick off the equivalent of a colony from your unknown plate. A visible inoculum should be seen on the tip and side of the wire.

2. Inoculate the EnteroPluri-Test by grasping the bent-end of the inoculating wire, twisting it, and withdrawing the wire through all 12 compartments using a turning motion.

3. Reinsert the wire into the tube (use a turning motion) through all 12 compartments until the notch on the wire is aligned with the opening of the tube. (The tip of the wire should be seen in the citrate compartment.) Break the wire at the notch by bending. Do not discard the wire yet.

4. Using the broken off part of the wire, punch holes through the cellophane which covers the air inlets located on the rounded side of the last 8 compartments. Your instructor will show you their correct location. Discard the broken off wire in the disinfectant container.

5. Replace both caps and incubate the EnteroPluri-Test on its flat surface at 36°- 37°C for 18-24 hours.

4. If your unknown is oxidase-positive,indicating a non-fermentative Gram-negative bacillus, do the following inoculation:

a. Streak your unknown for isolation on a plate of Cetrimide agar, a selective and differential medium for Pseudomonas, using one of the two streaking patterns illustrated in Fig. 4 and Fig. 5. Incubate upside down and stacked in the petri plate holder on the shelf of the 37°C incubator corresponding to your lab section.

Note that MacConkey agar can also be used to isolate Pseudomonas but we are using the Cetrimide agar today because it enables us to detect the production of the blue to green water-soluble pigment by Pseudomonas aeruginosa, as well as the production of fluorescein.

You will also inoculate an EnteroPluri-Test for practice only, but keep in mind that the EnteroPluri-Test is used to identify Enterobacteriaceae, not Pseudomonas.

Case Study #2

After receiving a baby chicken for Easter, a 7 year old boy is taken to the emergency room with symptoms of vomiting, nausea, non-bloody diarrhea, abdominal cramps, and a temperature of 100°F. A complete blood count (CBC) shows the WBC count to be within the reference range.

This XLD agar plate and this EnteroPluri-Test are from a stool culture from this patient.

CAUTION: TREAT THE UNKNOWN AS A PATHOGEN!. Inform your instructor of any spills or accidents. WASH AND SANITIZE YOUR HANDS WELL before leaving the lab.

Demonstration XLD agar plate and EnteroPluri-Test

PROCEDURE (to be done in groups of 3 )

1. Observe the following demonstrations shown in the links directly below and identify the causative bacterium:

a. An XLD agar plate,a selective medium used for isolating and differentiating Gram-negative enteric bacteria, especially intestinal pathogens such as Salmonella and Shigella.

2. Record your results in the Results section of Lab 13.

C. Lab Tests Used as Part of Today's Lab

To isolate Enterobacteriaceae and Pseudomonas, specimens from the infected site are plated out on any one of a large number of selective and differential media such as EMB agar, Endo agar, Deoxycholate agar, MacConkey agar, Hektoen Enteric agar, and XLD agar. We will look at three of these.

1. MacConkey Agar

MacConkey agar is a selective medium used for the isolation of non-fastidious Gram-negative rods, particularly members of the family Enterobacteriaceae and the genus Pseudomonas, and the differentiation of lactose fermenting from lactose non-fermenting Gram-negative bacilli. MacConkey agar contains the dye crystal violet well as bile salts that inhibit the growth of most Gram-positive bacteria but do not affect the growth of most Gram-negatives (see Fig. 6).

If the Gram-negative bacterium ferments the sugar lactose in the medium, the acid end products lower the pH of the medium. The neutral red in the agar turns red in color once the pH drops below 6.8. As the pH drops, the neutral red is absorbed by the bacteria, causing the colonies to appear bright pink to red.

  • Strong fementation of lactose with high levels of acid production by the bacteria causes the colonies and confluent growth to appear bright pink to red. The resulting acid, at high enough concentrations, can also causes the bile salts in the medium to precipitate out of solution causing a pink precipitate (cloudiness) to appear in the agar surrounding the growth(see Fig. 13).
  • Weak fermentation of lactose by the bacteria causes the colonies and confluent growth to appear pink to red, but without the precipitation of bile salts there is no pink halo around the growth(see Fig. 15).
  • If the bacteria do not ferment lactose, the colonies and confluent growth appear colorless and the agar surrounding the bacteria remains relatively transparent(see Fig. 17).

Typical colony morphology of our strains of Enterobacteriaceae and Pseudomonas aeruginosa on MacConkey agar is as follows:

1. Escherichia coli: colonies and confluent growth appear bright pink to red and surrounded by a pink precipitate (cloudiness) in the agar surrounding the growth (see Fig. 13). Strong fermentation of lactose.

2. Klebsiella pneumoniae: colonies and confluent growth appear bright pink to red but are not surrounded by a pink precipitate (cloudiness) in the agar surrounding the growth (see Fig. 14). Weak fermentation of lactose.

3. Enterobacter aerogenes: colonies and confluent growth appear bright pink to red but are not surrounded by a pink precipitate (cloudiness) in the agar surrounding the growth (see Fig. 15). Weak fermentation of lactose.

4. Enterobacter cloacae : colonies and confluent growth appear bright pink to red but are not surrounded by a pink precipitate (cloudiness) in the agar surrounding the growth (see Fig. 16). Weak fermentation of lactose.

5. Proteus mirabilis: colorless colonies agar relatively transparent (see Fig. 17). No fermentation of lactose.

6 . Proteus vulgaris: colorless colonies agar relatively transparent (see Fig. 18). No fermentation of lactose.

7 . Serratia marcescens : colorless colonies agar relatively transparent (see Fig. 19). No fermentation of lactose.

8 . Pseudomonas aeruginosa: colorless colonies agar relatively transparent (see Fig. 20).

9. Salmonella enterica: colorless colonies agar relatively transparent (see Fig. 21). No fermentation of lactose.

2 . XLD Agar

Xylose Lysine Desoxycholate (XLD) agar is used for isolating and differentiating Gram-negative enteric bacteria, especially intestinal pathogens such as Salmonella and Shigella. XLD agar contains sodium desoxycholate, which inhibits the growth of Gram-positive bacteria but permits the growth of Gram-negatives. It also contains the sugars lactose and sucrose, the amino acid L-lysine, sodium thiosulfate, and the pH indicator phenol red. Results can be interpreted as follows:

  • If the Gram-negative bacterium ferments lactose and/or sucrose, acid end products will be produced and cause the colonies and the phenol red in the agar around the colonies to turn yellow(see Fig. 16).
  • If lactose and sucrose are not fermented by the bacterium but the amino acid lysine is decarboxylated, ammonia, an alkaline end product will cause the phenol red in the agar around the colonies to turn a deeper red(see Fig. 17).
  • Sometimes the bacterium ferments the sugars producing acid end products and breaks down lysine producing alkaline end products. In this case some of the colonies and part of the agar turns yellow and some of the colonies and part of the agar turns a deeper red(see Fig. 18).
  • If hydrogen sulfide is produced by the bacterium as a result of thiosulfate reduction, part or all of the colony will appear black(see Fig. 19). Well-isolated colonies are usually needed for good results.

Typical colony morphology on XLD agar is as follows:

1. Escherichia coli: flat yellow colonies some strains may be inhibited.

2. Enterobacter and Klebsiella: mucoid yellow colonies.

3. Proteus: red to yellow colonies may have black centers.

4. Salmonella: usually red colonies with black centers.

5. Shigella, Serratia, and Pseudomonas: red colonies without black centers

Keep in mind, however, that some species and subspecies do not show typical reactions.

3. Cetrimide Agar (Pseudomonas P agar)

Cetrimide agar contains the chemical cetrimide (cetyl timethylammonium bromide) for the selective inhibition of most bacteria other than Pseudomonas. The medium also stimulates Pseudomonas aeruginosa to produce a number of water soluble iron chelators, including pyoverdin and pyocyanin. The green water soluble color characteristic of Pseudomonas aeruginosa is created when the yellow-green or yellow-brown fluorescent pyoverdin combines with the blue water-soluble pyocyanin (see Fig. 20). The fluorescent pyoverdin will typically fluoresce when the plate is placed under a short wavelength ultraviolet light (see Fig. 21). After a few minutes at room temperature, the plate loses its fluoresence. The fluoresence, however, can be restored by placing the plate back at 37°C for several minutes.

4. Oxidase Test

In this lab a Taxo N® disc is used to perform the oxidase test. The oxidase test is based on the bacterial production of an oxidase enzyme. Cytochrome oxidase, in the presence of oxygen, oxidizes the para-amino dimetheylanaline oxidase test reagent in a Taxo-N® disc.

  • In the immediate test, oxidase-positive reactions will turn a rose color within 30 seconds(see Fig. 5). Oxidase-negative will not turn a rose color (see Fig. 6). This reaction only lasts a couple of minutes.
  • In the delayed test, oxidase-positive colonies within 10 mm of the Taxo-N® disc will turn black within 20 minutes and will remain black(see Fig. 7). If the bacterium is oxidase-negative, the growth around the disc will not turn black (see Fig. 8).

Pseudomonas aeruginosa and most other non-fermentative, Gram-negative bacilli are oxidase-positive with the exception of the genus Plesiomonas, the Enterobacteriaceae are oxidase-negative.

5. Pigment production in Pseudomonas aeruginosa

The green water soluble color characteristic of Pseudomonas aeruginosa is created when the yellow-green or yellow-brown fluorescent pyoverdin combines with the blue water-soluble pyocyanin (see Fig. 20). The fluorescent pyoverdin will typically fluoresce when the plate is placed under a short wavelength ultraviolet light (see Fig. 21). After a few minutes at room temperature, the plate loses its fluoresence. The fluoresence, however, can be restored by placing the plate back at 37°C for several minutes. None of the Enterobacteriaceae produces pigment at 37°C.

6. Odor

Most of the Enterobacteriaceae have a rather foul smell Pseudomonas aeruginosa produces a characteristic fruity or grape juice-like aroma due to production of an aromatic compound called aminoacetophenone.

7. The EnteroPluri-Test

A number of techniques can be used for the identification of specific species and subspecies of Enterobacteriaceae. Speciation is important because it provides data regarding patterns of susceptibility to antimicrobial agents and changes that occur over a period of time. It is also essential for epidemiological studies such as determination of nosocomial infections and their spread.

In an effort to simplify the speciation of the Enterobacteriaceae and reduce the amount of prepared media and incubation space needed by the clinical lab, a number of self-contained multi-test systems have been commercially marketed. Some of these multi-test systems have been combined with a computer-prepared manual to provide identification based on the overall probability of occurrence for each of the biochemical reactions. In this way, a large number of biochemical tests can economically be performed in a short period of time, and the results can be accurately interpreted with relative ease and assurance.

The EnteroPluri-Test (see Fig. 22) is a self-contained, compartmented plastic tube containing 12 different agars (enabling the performance of a total of 15 standard biochemical tests) and an enclosed inoculating wire. After inoculation and incubation, the resulting combination of reactions, together with a Computer Coding and Identification System (CCIS), allows for easy identification. The various biochemical reactions of the EnteroPluri-Test and their correct interpretation are discussed below. Although it is designed to identify members of the bacterial family Enterobacteriaceae, it will sometimes also identify common biotypes of Pseudomonas and other non-fermentative Gram-negative bacilli. It does not identify Pseudomonas aeruginosa.

IDENTIFYING MEMBERS OF THE ENTEROBACTERIACEAE WITH THE ENTEROPLURI-TEST

The EnteroPluri-Test contains 12 different agars that can be used to carry out 15 standard biochemical tests (see Fig. 22). Interpret the results of your EnteroPluri-Test using the instructions below and record them on the EnteroPluri-Test table on your Results page. For more detail on the 15 biochemical tests in the EnteroPluri-Test, see Table 13A.

1. Interpret the results of glucose fermentation in compartment 1.

  • Any yellow = + red = -
  • If positive, circle the number 4 under glucose on your Results page.

2. Interpret the results of gas production also in compartment 1.

  • White wax lifted from the yellow agar = + wax not lifted from agar = -
  • If positive, circle the number 2 under gas on your Results page.

3. Interpret the results of lysine decarboxylase in compartment 2.

  • Any violet = + yellow = -
  • If positive, circle the number 1 under lysine on your Results page.

4. Interpret the results of ornithine decarboxylase in compartment 3.

  • Any violet = + yellow = -
  • If positive, circle the number 4 under ornithine on your Results page.

5. Interpret the results of H2S production in compartment 4.

  • Black/brown = + beige = - (The black may fade or revert back to negative if the EnteroPluri-Test is read after 24 hours of incubation.)
  • If positive, circle the number 2 under H2S on your Results page.

6. Indole production also in compartment 4. Do not interpret the indole test at this time. Add Kovac's reagent only after all other tests have been read (see step 16 below).

7. Interpret the results of adonitol fermentation in compartment 5.

  • Any yellow = + red = -
  • If positive, circle the number 4 under adonitol on your Results page.

8 . Interpret the results of lactose fermentation in compartment 6.

  • Any yellow = + red = -
  • If positive, circle the number 2 under lactose on your Results page.

9. Interpret the results of arabinose fermentation in compartment 7.

  • Any yellow = + red = -
  • If positive, circle the number 1 under arabinose on your Results page.

10. Interpret the results of sorbitol fermentation in compartment 8.

  • Any yellow = + red = -
  • If positive, circle the number 4 under sorbitol on your Results page.

11. Voges-Praskauer (VP) test in compartment 9. Do not interpret the VP test at this time. Add the reagents alpha-naphtol and potassium hydroxide (KOH) only after all other tests have been read (see step 17 below).

12. Interpret the results of dulcitol fermentation in compartment 10.

  • Yellow = + green or dark brown = -
  • If positive, circle the number 1 under dulcitol on your Results page.

13. Interpret the results of PA deaminase also in compartment 10.

  • Dark brown= + green or yellow= -
  • If positive, circle the number 4 under PA on your Results page.

14. Interpret the results of urea hydrolysis in compartment 11.

  • Pink, red or purple = + beige = -
  • If positive, circle the number 2 under urea on your Results page.

15. Interpret the results of citrate utilization in compartment 12.

  • Any blue = + green = -
  • If positive, circle the number 1 under citrate on your Results page.

16. Your instructor will add 2-3 drops of Kovac's reagent to the indole test compartment.

17. Your instructor will add 3 drops of alpha-naphtol reagent and 2 drops of potassium hydroxide (KOH) to the VP test compartment.

18. Add all the positive test number values in each bracketed section and enter each sum in its code box on the EnteroPluri-Test chart on your Results page.

19. The 5 digit number is the CODICE number. Look that number up in the Codebook and identify your unknown. (Should more than one organism be listed, the confirmatory tests indicated in the CCIS would normally then have to be performed. In addition, an identification of Salmonella or Shigella would usually be confirmed by direct serologic testing as will be described in Lab 17.)


Regulation of acetate metabolism in Escherichia coli BL21 by protein N ε -lysine acetylation

Acetate production is one of the most striking differences between Escherichia coli K12 and BL21 strains. Transcription of acetate metabolism genes is regulated. Additionally, acetyl-CoA synthetase, which activates acetate to acetyl-CoA, is regulated by post-translational acetylation. The aim of this study was to understand the contribution of reversible protein lysine acetylation to the regulation of acetate metabolism in E. coli BL21. The phenotypic differences between both strains were especially important in the presence of acetate. The high expression of acetyl-CoA synthetase (acs) in glucose exponential phase in BL21 allows the simultaneous consumption of acetate and glucose. Lack of catabolite repression also affected its post-translational regulator, the protein acetyltransferase (patZ). The effect of the deletion of cobB (encoding a sirtuin-like protein deacetylase) and patZ genes depended on the genetic background. The deletion of cobB in both strains increased acetate production and decreased growth rate in acetate cultures. The deletion of patZ in BL21 suppressed acetate overflow in glucose medium and increased the growth rate in acetate cultures. Differences on acetate overflow between BL21 and K12 strains are caused by many overlapping factors. Two major contributing effects were identified: (1) the expression of acs during exponential growth is not repressed in the BL21 strain due to concomitant cAMP production and (2) the acetyl-CoA synthetase activity is more tightly regulated by protein acetylation in BL21 than in the K12. Altogether these differences contribute to the lower acetate overflow and the improved ability of E. coli BL21 to consume this metabolite in the presence of glucose.

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Fate of bacterial biomass derived fatty acids in soil and their contribution to soil organic matter

Soil organic matter (SOM) is a major pool of the global C cycle and determines soil fertility. The stability of SOM strongly depends on the molecular precursors and structures. Plant residues have been regarded as the dominant precursors, but recent results showed a major contribution of microbial biomass. The fate of microbial biomass constituents has not yet been explored therefore, we investigated the fate of fatty acids (FA) from 13 C labeled Gram-negative bacteria (Escherichia coli) in a model soil study [Kindler, R., Miltner, A., Richnow, H.H., Kästner, M., 2006. Fate of gram negative bacterial biomass in soil—mineralization and contribution to SOM. Soil Biology & Biochemistry 38, 2860–2870]. After 224 days of incubation, the label in the total fatty acids (t-FA) in the soil decreased to 24% and in the phospholipid fatty acids (PLFA) of living microbes to 11% of the initially added amount. Since the bulk C decreased only to 44% in this period, the turnover of FA is clearly higher indicating that other compounds must have a lower turnover. The 13 C label in the t-FA reached a stable level after 50 days but the label of the PLFA of the living microbial biomass declined until the end of the experiment. The isotopic enrichment of individual PLFA shows that the biomass derived C was spread across the microbial food web. Modelling of the C fluxes in this experiment indicated that microbial biomass is continuously mineralized after cell death and recycled by other organisms down to the 10% level, whereas the majority of biomass derived residual bulk C (∼33%) was stabilized in the non-living SOM pool.

Present address: Department of Waste Management and Environmental Research, Berlin University of Technology, Franklinstr. 29, 10587 Berlin, Germany.


Metabolism

2. the sum of the physical and chemical processes by which living organized substance is built up and maintained ( anabolism ), and by which large molecules are broken down into smaller molecules to make energy available to the organism ( catabolism ). Essentially these processes are concerned with the disposition of the nutrients absorbed into the blood following digestion.

There are two phases of metabolism: the anabolic and the catabolic phases. The anabolic, or constructive, phase is concerned with the conversion of simpler compounds derived from the nutrients into living, organized substances that the body cells can use. In the catabolic, or destructive, phase these organized substances are reconverted into simpler compounds, with the release of energy necessary for the proper functioning of the body cells.

The rate of metabolism can be increased by exercise by elevated body temperature, as in a high fever, which can more than double the metabolic rate by hormonal activity, such as that of thyroxine, insulin, and epinephrine and by specific dynamic action that occurs following the ingestion of a meal.

The basal metabolic rate refers to the lowest rate obtained while an individual is at complete physical and mental rest. Metabolic rate usually is expressed in terms of the amount of heat liberated during the chemical reactions of metabolism. About 25 per cent of all energy from nutrients is utilized by the body to carry on its normal function the remainder becomes heat.


Effects of Paracetamol on NOS, COX, and CYP Activity and on Oxidative Stress in Healthy Male Subjects, Rat Hepatocytes, and Recombinant NOS

Paracetamol (acetaminophen) is a widely used analgesic drug. It interacts with various enzyme families including cytochrome P450 (CYP), cyclooxygenase (COX), and nitric oxide synthase (NOS), and this interplay may produce reactive oxygen species (ROS). We investigated the effects of paracetamol on prostacyclin, thromboxane, nitric oxide (NO), and oxidative stress in four male subjects who received a single 3 g oral dose of paracetamol. Thromboxane and prostacyclin synthesis was assessed by measuring their major urinary metabolites 2,3-dinor-thromboxane B2 and 2,3-dinor-6-ketoprostaglandin F1α, respectively. Endothelial NO synthesis was assessed by measuring nitrite in plasma. Urinary 15(S)-8-iso-prostaglanding F2α was measured to assess oxidative stress. Plasma oleic acid oxide (cis-EpOA) was measured as a marker of cytochrome P450 activity. Upon paracetamol administration, prostacyclin synthesis was strongly inhibited, while NO synthesis increased and thromboxane synthesis remained almost unchanged. Paracetamol may shift the COX-dependent vasodilatation/vasoconstriction balance at the cost of vasodilatation. This effect may be antagonized by increasing endothelial NO synthesis. High-dosed paracetamol did not increase oxidative stress. At pharmacologically relevant concentrations, paracetamol did not affect NO synthesis/bioavailability by recombinant human endothelial NOS or inducible NOS in rat hepatocytes. We conclude that paracetamol does not increase oxidative stress in humans.

1. Introduction

Nitric oxide (NO), prostaglandin (PG) I2, that is, prostacyclin (PGI2), and thromboxane A2 (TxA2) are important short-lived signaling molecules involved in many physiological and pathological processes. Thus, PGI2 and NO are potent vasodilators and inhibitors of platelet aggregation. Contrarily, TxA2 is a strong vasoconstrictor and inductor of platelet aggregation. NO is synthesized from L-arginine (Arg) by constitutive and inducible NO synthase (NOS) isoforms. Prostaglandin H synthase (PGHS) isoforms, generally termed cyclooxygenase (COX), convert arachidonic acid (AA) to the collectively named prostanoids. The L-arginine/NO pathway is generally accepted to interact with the COX pathway and to modulate its activity [1–4]. For instance, the inducible NOS (iNOS) isoform has been shown to bind to the inducible COX isoform (COX-2) and to

-nitrosylate and activate COX-2 [2]. The role of NO in prostaglandin biology has been recently updated by Kim [4]. Potential mechanisms of direct NOS-COX cross-talk may include (1) binding of NO to the iron atom of the heme group of COX, (2) reaction of the nitrosyl cation (NO + ) with sulfhydryl (SH) groups of cysteine (Cys) moieties of COX to form -nitroso-COX, and (3) reaction of peroxynitrite (ONOO − ), that is, the reaction product of NO radical ( NO) and superoxide radical anion (

) produced either by NOS itself or by other enzymes including COX and CYP [3], with SH groups of Cys residues or with tyrosine (Tyr) residues of COX being involved in the catalytic process [2]. -Nitrosylation of COX-Cys moieties by higher oxides of NO, notably dinitrogen trioxide (N2O3), and by ONOO − and -transnitrosylation of COX-Cys moieties by low-molecular-mass -nitrosothiols have been shown to both enhance and inhibit COX activity. Nitration of Tyr residues located in the catalytic domain of COX is assumed to inhibit COX activity [2, 4–6]. On the other hand, ONOO − has been reported to enhance COX activity presumably by increasing the peroxide concentration that is required for the peroxidase activity of COX [7].

Paracetamol (acetaminophen, APAP) is one of the most frequently applied drugs worldwide and is considered generally a safe analgesic and antipyretic drug in therapeutic dosage, which lacks however appreciable anti-inflammatory and antiplatelet activity [10]. The mechanism of the analgesic and antipyretic effects of paracetamol is not fully established, yet inhibition of PGHS activity by paracetamol in different cell and tissue types is generally assumed to be the main mode of paracetamol’s analgesic and antipyretic action. PGHS possesses both peroxidase and cyclooxygenase activity. Paracetamol is believed to inhibit the peroxidase catalytic site of PGHS, unlike the majority of nonsteroidal anti-inflammatory drugs (NSAIDs) and the PGHS2 inhibitors. In vitro, paracetamol is a much stronger inhibitor of prostanoid synthesis in endothelial cells than in platelets. In particular, paracetamol is a weak inhibitor of TxA2 synthesis in platelets. It is also remarkable that the inhibitory potency of paracetamol is inversely correlated with the PGHS concentration (for a review, see [10]). These particular characteristics distinguish paracetamol from NSAIDs including acetylsalicylic acid (ASA).

In vitro, PGHS activity can be assessed by measuring the production rate of various primary prostanoids, such as PGE2, PGI2, and TxA2. Because of the remarkable chemical instability of PGI2 and TxA2, their stable hydrolysis products, that is, 6-keto-PGF1α and TxB2, respectively, are measured instead of PGI2 and TxA2 [11]. In vivo, measurement of PGE2, 6-keto-PGF1α, and TxB2 in plasma is associated with artefactual prostanoid synthesis during blood sampling and may lead to incorrect conclusions with regard to PGHS activity [12]. This especially applies to TxA2 which is produced in high amounts in activated platelets [13]. Measurement of PGE2 in the urine reflects renal PGE2 synthesis. By far more reliable is the measurement of major urinary metabolites of prostanoids, such as 2,3-dinor-TxB2 for TxA2, 2,3-dinor-6-keto-PGF1α for PGI2, and the major urinary metabolite of PGE2 (PGE-MUM) for systemic PGE2 production [11]. This can be best accomplished by means of analytical technologies which have high inherent sensitivity and selectivity such as gas chromatography-mass spectrometry (GC-MS) and more so gas chromatography-tandem mass spectrometry (GC-MS/MS) (for a review, see [11]).

Recently, Sudano and colleagues [9] reported that paracetamol (1 g TID for 2 weeks on top of standard cardiovascular therapy) increased ambulatory mean systolic and diastolic blood pressure by about 3 and 2 mmHg, respectively, without changing endothelium and platelet function in patients with coronary artery disease (CAD). Sudano et al. [9] concluded that, particularly in patients at increased cardiovascular risk, use of paracetamol should be evaluated as rigorously as traditional NSAIDs and selective COX2 inhibitors. In that study, plasma and urine PGE2 as well as plasma TxB2 did not change upon paracetamol administration [9]. However, as mentioned above, measurement of PGE2 and TxB2 in plasma is prone to artefactual prostanoid synthesis [12, 13], whereas measurement of PGE2 in the urine does not provide information about PGHS-catalyzed synthesis of the two antagonists TxA2 and PGI2 [11].

In humans, oral administration of paracetamol (500 mg) has been reported not to result in decreased excretion rate of 2,3-dinor-TxB2, unlike aspirin (500 mg) or indomethacin (50 mg), as measured by GC-MS [14]. Also, in contrast to aspirin (3 g for 2 days), oral administration of paracetamol (3 g for 2 days) has been reported not to reduce urinary excretion of PGE2 but to weakly reduce PGE-MUM excretion indicating inhibition of systemic PGE2 synthesis [15]. On the other hand, a single oral dose of 500 mg paracetamol has been shown to reduce urinary excretion rate of 2,3-dinor-6-keto-PGF1α for 6–8 h by maximally 60% (i.e., inhibition of PGI2 synthesis), without reducing urinary excretion rate of 2,3-dinor-TxB2 (i.e., no inhibition of TxA2 synthesis) [16]. The results of these in vivo studies in human subjects suggest that orally administered paracetamol, at a single dose of 500 mg or at a cumulative dose of 3000 mg per day, does not inhibit remarkably TxA2 synthesis, but it may temporarily inhibit PGI2 synthesis.

The ramifications between NOS and COX pathways have been frequently investigated in the past (reviewed in [4]), but results are inconsistent. For instance, in murine macrophages, paracetamol, at pharmacologically relevant plasma concentrations (60–120 μM), has been reported not to affect iNOS activity [17]. At suprapharmacological concentrations (2, 5, and 10 mM), paracetamol has been reported to inhibit iNOS gene expression and iNOS activity in RAW 264.7 cell line macrophages [18]. By contrast, paracetamol (up to 10 mM) has been reported not to affect neuronal NOS (nNOS) and iNOS activity in rat cerebellum and HUVECs [19]. Others have reported that paracetamol (100 μM) did not affect nNOS activity in cerebellum but inhibited NOS activity in murine spinal cord slices as measured by the radiolabelled L-citrulline assay [20]. The effect of paracetamol on in vivo in humans is elusive.

Because paracetamol, when applied at pharmacological doses, inhibits the synthesis of the vasodilatatory and antiaggregatory PGI2 much stronger and sustainably than the synthesis of the vasoconstrictory and thrombogenic TxA2 in humans, we wondered whether the paracetamol-induced shifting of the balance between vasodilatatory/antiaggregatory and vasoconstrictory/thrombogenic COX-related homeostasis may induce processes that lead to enhanced synthesis of the vasodilatatory/antiaggregatory NO, thus counteracting blood pressure fall and platelet activation. Preliminary investigations of our group showed that paracetamol, administered in therapeutic doses to healthy humans (up to 10 mg/kg), did not change whole body NO synthesis (data not shown), suggesting that a potential effect of paracetamol on NOS activity is likely to require much higher, suprapharmacological doses of this drug. In consideration of the toxicological potency of high paracetamol doses, we investigated the effects of a single oral 3 g dose in four healthy volunteers. To our knowledge, the effects of such a high single oral dose of paracetamol on PGI2, TxA2, and NO synthesis in humans have not been investigated so far. Because of the high dose used in the human study, paracetamol may induce oxidative stress and decrease NO bioavailability [21]. We therefore measured the oxidative stress biomarker 15(S)-8-iso-PGF2α [22] in plasma and urine. Nitrite in plasma was measured as a biomarker of NO synthesis and bioavailability (reviewed in [23]). In addition, we performed in vitro studies on recombinant endothelial NOS (eNOS) and inducible NOS (iNOS) in rat hepatocytes to test potential effects of paracetamol on NO synthesis and bioavailability.

2. Materials and Methods

2.1. Subjects and Study Performance

Four healthy nonsmoking male adults (aged 39, 40, 44, and 64 years) participated in the study and gave their informed consent to the study. The volunteers received orally six 500 mg paracetamol tablets (Ratiopharm) at once. Dosage was each 29 mg/kg for volunteer A and volunteer B, 37 mg/kg for volunteer C, and 52 mg/kg for volunteer D. Volunteers were not fasting but they did not eat in the first three hours following paracetamol administration. Before and after paracetamol administration, venous blood and urine were collected in 30 and 60 min intervals over an observation period of 6 h for analysis of biochemical parameters as described below. Venous blood (8 mL) was drawn by using 9 mL EDTA vacutainers (Sarstedt, Germany) and centrifuged immediately (800 ×g, 4°C, 5 min). Plasma was decanted, portioned in 0.1 and 1.0 mL aliquots as required for each biochemical parameter, and stored frozen at −80°C until analysis. Urine from spontaneous micturition was collected in 45 mL polypropylene tubes, aliquoted in 0.1 and 1.0 mL portions according to the requirement of the individual biochemical parameters, and stored at −20°C until analysis.

2.2. Analysis of Biochemical Parameters in the Human Study

All samples of this study were analyzed within 10 days after collection. In GC-MS and GC-MS/MS methods, stable-isotope labelled analogs were used as internal standards as reported in the respective references cited below. We found that paracetamol added to pooled human plasma at concentrations of 10, 25, 50, 75, and 100 mg/L did not interfere with the analysis of the biochemical parameters measured in the study plasma samples (data not shown). Data from this study are reported as mean ± standard error of the mean (SEM).

2.2.1. Measurement of Paracetamol

Plasma paracetamol concentration was determined by reverse phase HPLC (250 × 4 mm i.d., 5 μm particle size) with isocratic elution (mobile phase: 45 mM ammonium sulphate-acetonitrile, 10 : 1, v/v flow rate: 1 mL/min) with UV absorbance detection at 236 nm.

2.2.2. Measurement of Prostanoids and Creatinine

PGI2 and TxA2 synthesis was assessed by GC-MS/MS by measuring in 1 mL urine aliquots the respective major urinary metabolites [11], that is, 2,3-dinor-6-keto-PGF1α and 2,3-dinor-TxB2, exactly as described elsewhere [24]. PGE2 and free nonconjugated 15( )-8-iso-PGF2α in urine (1 mL) and free 15(S)-8-iso-PGF2α in plasma (1 mL) were measured by GC-MS/MS after extraction by immunoaffinity column chromatography as described previously [25]. Urinary excretion rate of the eicosanoids was corrected for creatinine excretion [11] and is expressed in nmol prostanoid/mol creatinine. Urine creatinine was measured in 10 μL urine aliquots by GC-MS as reported elsewhere [26].

2.2.3. Analysis of the L-Arginine/NO Pathway

Nitrite and nitrate were measured simultaneously in 100 μL aliquots of plasma or urine by GC-MS as described elsewhere [27]. Urinary excretion rate of nitrite and nitrate was corrected for creatinine excretion as well. Arginine and the endogenous NOS activity inhibitor asymmetric dimethylarginine (ADMA) were measured by GC-MS and GC-MS/MS, respectively, in 100 μL aliquots of ultrafiltrate obtained from plasma by centrifugation according to previously reported procedures [8].

2.2.4. Additional Analyses

Total homocysteine (hCys) in plasma (0.1 mL) was measured by a commercially available fluorescence polarimetry immunoassay (FPIA). In vivo CYP activity [28] was assessed by measuring oleic acid oxide (cis-EpOA) in 1 mL aliquots of plasma as described elsewhere [29].

2.2.5. Quality Control

Quality control (QC) samples were analyzed alongside study samples for all biochemical parameters. Accuracy and precision in the QC samples were within generally accepted ranges that is, bias and imprecision levels were below 20%.

2.3. Effect of Paracetamol on Recombinant Human eNOS Activity

The effect of paracetamol on NOS activity in vitro was investigated by using a commercially available (ALEXIS, Grünberg, Germany) recombinant human endothelial NOS (heNOS) and by measuring simultaneously formation of [ 15 N]nitrite and [ 15 N]nitrate from L-[guanidine- 15 N2]arginine by means of a GC-MS assay [30]. Incubations were performed at 37°C in 50 mM potassium phosphate buffer (1000 μL, pH 7) containing heNOS (50 μg/mL), L-[guanidine- 15 N2]-arginine (20 μM, Cambridge Isotope Labs, Andover, MA, USA), and all NOS cofactors (all purchased from Sigma-Aldrich, Steinheim, Germany) and prosthetic groups (10 μM tetrahydrobiopterin, 800 μM NADPH, 5 μM FAD, 5 μM for FMN, 500 nM calmodulin, and 500 μM CaCl2 (Merck, Darmstadt, Germany)). Reactions were terminated by addition of 400 μL aliquots of ice cold acetone and samples were processed for GC-MS analysis of [ 15 N]nitrite and [ 15 N]nitrate. Unlabeled nitrite and nitrate were used as internal standards for [ 15 N]nitrite and [ 15 N]nitrate, respectively. Data are shown as mean ± standard deviation (SD) from two independent experiments.

2.4. Effect of Paracetamol on iNOS Activity in Cultured Rat Hepatocytes Proliferating In Vitro

The effect of paracetamol on iNOS activity was investigated in primary rat hepatocytes proliferating in vitro as described recently by measuring formation of [ 15 N]nitrite and [ 15 N]nitrate by GC-MS [31]. In some experiments, LiCl (10 mM) was used to enhance expression of iNOS-mRNA and cell growth [31]. Incubations were performed at 37°C in the presence of 5 mM L-[guanidine- 15 N2]-arginine added at the time -22 h. Reactions were terminated by addition of 400 μL aliquots of ice cold acetone, and samples were processed for GC-MS analysis of [ 15 N]nitrite and [ 15 N]nitrate. Data are shown as mean ± SD from three independent experiments.

2.5. Statistical Analysis

Because of considerable differences in the baseline concentrations of some of the biochemical parameters measured in the four subjects, changes and statistical significance were calculated by setting the respective baseline levels to 100%. Statistical significance (

) was evaluated by using unpaired

-test and comparing the data obtained at various times to the baseline values or to the 0.5 h values when percentage changes were compared.

3. Results

3.1. Effect of High-Dose Paracetamol on COX, NOS, and CYP Activity and on Oxidative Stress in Humans

Mean maximum paracetamol plasma concentration (

) was 30.2 mg/L (200 μmol/L) (Figure 1). This value is consistent with a value of about 20 mg/L that has been reached after oral administration of 2000 mg of paracetamol [32]. In the urine samples, we measured by reverse phase HPLC with UV absorbance detection comparable creatinine-corrected excretion rates of the paracetamol glucuronide and sulphate metabolites (data not shown).


Plasma concentration of paracetamol before and after administration (time zero is indicated by the dashed arrow) of a single oral 3 g dose of paracetamol to four male subjects. Data are shown as mean ± SEM.

Upon 3 g paracetamol intake, considerable and sustained decrease in creatinine-corrected 2,3-dinor-6-keto-PGF1α excretion rate was seen, suggesting strong PGI2 inhibition by paracetamol (Figure 2(a)). Maximum and statistically significant PGI2 inhibition of about 60 to 70% was reached 1 h, 1.5 h, and 2.5 h after paracetamol administration. The extent of the decrease seen in the 2,3-dinor-6-keto-PGF1α excretion rate in the present study is comparable to that seen upon administration of a 500 mg oral paracetamol dose [14]. Paracetamol caused only a moderate, statistically insignificant decrease in 2,3-dinor-TxB2 excretion in the four volunteers (Figure 2(b)). In three out of the four volunteers, maximum TxA2 inhibition of about 70% was reached 1.5 h after administration, but the duration of TxA2 inhibition was relatively short (not shown). Figure 2(c) shows that the PGI2/TxA2 molar ratio decreased by a factor of 2 to 3 upon paracetamol administration, although statistical significance failed by a hair 1.5 h (

) and 2.5 h ( ) after paracetamol ingestion. Paracetamol seemed to decrease very weakly the excretion of PGE2 in the urine (Figure 2(d)), suggesting that even 3 g of paracetamol taken at once is not able to inhibit renal synthesis of PGE2 in the four volunteers enrolled in the study.


(a)
(b)
(c)
(d)
(a)
(b)
(c)
(d) Effect of a single oral 3 g dose of paracetamol on systemic prostacyclin and thromboxane synthesis and on renal synthesis of PGE2 in four healthy volunteers (time zero and baseline values are indicated by dashed arrows). (a) Creatinine-corrected urinary excretion of 2,3-dinor-6-keto-prostaglandin F1α (2,3-dn-6k-PGF1α) as a measure of systemic PGI2 synthesis. (b) Creatinine-corrected urinary excretion of 2,3-dinor-thromboxane B2 (2,3-dn-TxB2) as a measure of systemic TxA2 synthesis. (c) PGI2/TxA2 molar ratio calculated from the 2,3-dn-6k-PGF1α and 2,3-dn-TxB2 excretion rates shown in (a) and (b), respectively. (d) Creatinine-corrected urinary excretion of PGE2 as a measure of renal PGE2 synthesis. An asterisk in (a) indicates statistical significance (

Paracetamol-induced changes in systemic prostacyclin synthesis were not accompanied by noteworthy changes in the plasma concentration of total hCys (Figure 3(a)) or in the urinary excretion rate of free 15(S)-8-iso-PGF2α (Figure 3(b)), suggesting no elevation or reduction of oxidative stress upon high-dosed paracetamol administration.


(a)
(b)
(a)
(b) Effect of a 3 g single oral dose of paracetamol on oxidative stress in four healthy volunteers (time zero and baseline value are indicated by dashed arrows). (a) Plasma total homocysteine (hCys) concentration and (b) creatinine-corrected urinary excretion rate of 15(S)-8-iso-prostaglandin F2α (15(

Because of the considerable difference in the baseline plasma nitrite and nitrate concentrations measured in the four subjects, changes in plasma nitrite and nitrate were calculated and presented as percentage of the respective baseline levels. Figure 4(a) shows moderate increases in plasma nitrite concentration which were statistically significantly higher when the 2.5 and 3.5 h values were compared with the 0.5 h values. Changes in plasma nitrate concentrations (Figure 4(b)) and urine nitrite (Figure 4(c)) and urine nitrate (Figure 4(d)) excretion were not statistically significantly different. Finally, plasma Arg and ADMA concentrations did not change upon paracetamol ingestion (Figure 4(e)).


(a)
(b)
(c)
(d)
(e)
(a)
(b)
(c)
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(e) Effect of a 3 g single oral dose of paracetamol on plasma (a) and urine (c) nitrite, plasma (b) and urine (d) nitrate, and plasma arginine (Arg) and asymmetric dimethylarginine (ADMA) (e) in four healthy volunteers (time zero and baseline values are indicated by dashed lines). Data in plasma are shown as percentage changes of the baseline plasma nitrite concentrations (1.26, 3.04, 3.76, and 4.05 μM) and baseline plasma nitrate concentrations (37.3, 42.5, 26.8, and 52.9 μM), respectively. Data are shown as mean ± SEM. An asterisk indicates statistical significance (

Previously, we showed that the whole body activity of CYP isoforms can be assessed by measuring the concentration of the free, that is, nonesterified, oleic acid oxide cis-EpOA in plasma [28]. Figure 5 shows an abrupt increase in mean plasma cis-EpOA concentration 2.5 h after paracetamol administration followed by an abrupt fall to baseline level 1 h later. This finding may suggest a very short-term paracetamol-induced elevation of CYP activity. However, we also found a very similar change in the plasma concentration of free 15( )-8-iso-PGF2α (Figure 5). Previously, we observed that addition of phospholipase A2 (PLA2) to human serum increased in parallel the concentration of both free cis-EpOA and free 15( )-8-iso-PGF2α [25]. Therefore, the temporary short-time increases in cis-EpOA and 15(S)-8-iso-PGF2α seen 2.5 h after paracetamol administration may have resulted from release of presumably hepatic PLA2 into the blood.


Effect of a 3 g single oral dose of paracetamol on serum cis-epoxyoctadecanoic acid (cis-EpOA) and 15(

)-8-iso-PGF2α in four healthy volunteers (time zero and baseline value are indicated by dashed lines). Data are shown as mean ± SEM. Note the logarithmic scale on the

-axis. Only the 2.5 h concentration of cis-EpOA was statistically significantly different (

3.2. Effects of Paracetamol on Recombinant heNOS Activity and iNOS in Rat Hepatocytes

At the therapeutically relevant concentration of 100 μM (i.e., 15 mg/L), paracetamol had only a weak effect on the formation of [ 15 N]nitrite and [ 15 N]nitrate in incubation mixtures of a recombinant heNOS (Figure 6). Linear regression of analysis between the concentrations of [ 15 N]nitrite and [ 15 N]nitrate in the presence of paracetamol versus the concentrations of [ 15 N]nitrite and [ 15 N]nitrate in the absence of paracetamol revealed a slope value of 0.834 (Figure 6(b)). This finding suggests that paracetamol inhibited heNOS-catalyzed 15 NO formation (i.e., [ 15 N]nitrite and [ 15 N]nitrate) from L-[guanidine- 15 N2]-arginine in average by 16.6%, notably for incubation times longer than 10 min. Similar small effects of paracetamol on iNOS were also seen in experiments with adult rat hepatocytes proliferating in vitro independent of the presence of LiCl (Figure 7). It is well established that peroxynitrite can nitrate paracetamol to 3-nitroparacetamol [33]. In the paracetamol-containing samples from both the recombinant heNOS and the rat hepatocytes iNOS, no 3-[ 15 N]nitroparacetamol was detected by GC-MS/MS above the limit of quantitation (about 1 nM), suggesting no formation of peroxynitrite (data not shown).


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(b) Effect of paracetamol (APAP) at 100 μM (15 mg/L) on the formation of [ 15 N]nitrite and [ 15 N]nitrate in an incubation mixture of recombinant heNOS upon incubation time (a) and linear regression analysis between [ 15 N]nitrite and [ 15 N]nitrate concentrations measured in the presence and absence of paracetamol (b). Data in (a) are shown as mean ± SD from two independent experiments no statistical analysis was performed.

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(b) Effect of paracetamol (APAP) at 100 μM (15 mg/L) on the peak area ratio of m/z 47 for [ 15 N]nitrite to m/z 46 for [ 14 N]nitrite (a) and on the peak area ratio of m/z 63 for [ 15 N]nitrate to m/z 62 for [ 14 N]nitrate (b) upon incubation of adult rat hepatocytes with L-[guanidine- 15 N2]-arginine (5 mM) in the absence and in the presence of LiCl (1 mM) for the indicated times at 37°C as described elsewhere [8]. Reactions were terminated by addition of 400 μL aliquots of ice cold acetone and samples were further processed for GC-MS analysis. Data are shown as mean ± SD from three independent experiments no statistical analysis was performed.

4. Discussion

4.1. General Remarks and Aim of the Study

Paracetamol is generally assumed to increase oxidative stress and is therefore commonly used in animal models of oxidative stress, in which paracetamol is administered in exorbitant high doses [21]. Whether paracetamol, when administered at therapeutic doses, also acts as a prooxidant is unknown. Paracetamol is known to interact with many enzymes such as CYP, COX, and NOS, which themselves are known to contribute to oxidative stress, for instance, by producing superoxide radical anions. While the inhibitory effect of paracetamol on prostacyclin synthesis in vivo in humans is well established [16], its effects on thromboxane and NO synthesis as well as on CYP activity are incompletely understood. This may be due to insufficiently high intracellular paracetamol concentrations when this drug is administered in therapeutic doses, for instance, by oral administration of a 500 mg paracetamol tablet. The aim of the present work was to investigate in healthy humans the effects of high-dosed paracetamol (i.e., 3 g) on the activity of the COX, NOS, and CYP, as well as on oxidative stress. Given the well-known hepatotoxicity of paracetamol, only four healthy subjects were enrolled in the human study. By using paracetamol concentrations that are expected to prevail for a considerable period of time after administration of a single 3 g oral dose to humans, we investigated the effects of paracetamol at suprapharmacological concentrations on the activity of two NOS isoforms in vitro, that is, on recombinant human eNOS and iNOS in rat hepatocytes.

4.2. Effects of Paracetamol on the Cyclooxygenase Pathway

Considering a mean fraction (oral bioavailability,

) value of 88% for paracetamol [32], its mean distribution volume (

) in the volunteers of the human study described in this paper is estimated to be 88 L. This value is almost 25 times higher than the estimated volunteers’ plasma volume and suggests that paracetamol may reach concentrations up to about 5000 μM in other body compartments except for red blood cells. Such high concentrations would be high enough to inhibit prostacyclin (PGI2) and thromboxane (TxA2) synthesis in endothelial cells and platelets, respectively [10].

Indeed, paracetamol, at the high single oral dose of 3 g, potently inhibited PGHS-catalyzed synthesis of PGI2, a potent vasodilator and inhibitor of platelet aggregation. By contrast, the synthesis of TxA2, a potent vasoconstrictor and platelet activator, was found not to be significantly inhibited by paracetamol in four subjects. Relative effects of drugs on PGI2 and TxA2 synthesis are commonly estimated by using the molar ratio of the prostanoids [34]. In our study, oral administration of a single 3 g oral paracetamol dose decreased the average PGI2/TxA2 molar ratio from about 0.6 before administration to values ranging between 0.4 and 0.2 after administration, thus shifting the vasodilatatory (PGI2)/vasoconstrictory (TxA2) balance at the cost of vasodilatation. A consequence of such a shift may be an increase of blood pressure. Indeed, Sudano and colleagues found that chronic administration of paracetamol to CAD patients at a lower dose (1 g TID for 2 weeks) than in the present study resulted in small blood pressure increase [9]. It is worth mentioning that in the study by Sudano et al. [9] considerably lower plasma paracetamol concentrations had been reached in comparison to those we measured in our study. In confirmation of previous studies [9, 15], we found that excretion of PGE2 did not change upon paracetamol administration, suggesting that even a high dose of 3 g paracetamol did not alter significantly renal PGE2 production in the volunteers.

4.3. Effects of Paracetamol on the L-Arginine/NO Pathway

The effects of paracetamol on NOS expression and activity have been studied by several groups. Yet, the observations are contradictory [17–20]. In our human study, paracetamol increased temporarily the concentration of nitrite in plasma. As the major fraction of circulating nitrite may originate from NO produced in the endothelium [23], our in vivo results may indicate that paracetamol increased eNOS activity and/or eNOS expression 2.5 to 3.5 h after administration. Yet, alternative ways such as paracetamol-induced reduction of nitrate to nitrite may have also increased plasma nitrite concentrations in the volunteers. Paracetamol did not change the plasma concentration of two other main parameters of the L-arginine/NO pathway, that is, L-arginine and ADMA. In vitro, paracetamol had only a very weak inhibitory effect on isolated recombinant heNOS and on iNOS in adult rat hepatocytes proliferating in vitro. LiCl that is known to induce expression and activity of iNOS in rat hepatocytes [31] increased iNOS activity but did not alter NO bioavailability. Thus, neither paracetamol nor LiCl influenced iNOS-related oxidative stress in rat hepatocytes.

4.4. Effects of Paracetamol on the Cytochrome P450 Pathway

Paracetamol is oxidized by the CYP family to NAPQI (

-benzoquinone imine), the toxic intermediate of paracetamol. Unsaturated fatty acids including arachidonic acid and oleic acid are substrates for CYP enzymes [28, 35], and some of the arachidonic acid epoxides are vasoactive compounds [35]. At high concentrations (e.g., 1000 μM), paracetamol may inhibit the activity of CYP isoforms. In our human study, paracetamol increased temporarily the plasma concentration of the oleic acid oxide cis-EpOA. As cis-EpOA is a marker of CYP activity in humans [28], this finding may suggest that paracetamol increases CYP activity for a very short period of time. Another explanation for the very short-lasting increase in plasma cis-EpOA concentration could be activation of extracellular phospholipase A2 (PLA2) activity or release of hepatic PLA2 into the blood stream by paracetamol, because a considerable fraction of cis-EpOA is found esterified to human serum lipids [28]. The latter explanation is supported by the finding that the plasma concentration of free 15( )-8-iso-PGF2α displayed a similar course including the sharp maximum like cis-EpOA in the present human study. It is worth mentioning that both 15(S)-8-iso-PGF2α and cis-EpOA are released in parallel from serum lipids upon incubation with PLA2 [28]. At present, very little is known about paracetamol effects on PLA2 activity and/or expression. In contrast to indomethacin, paracetamol (at 1000 μM) was found not to inhibit extracellular PLA2 activity as measured using radiolabelled oleic acid esterified to E. coli membranes [36]. In mice, hepatotoxicity induced by paracetamol at a dose of 400 mg/kg, that is, about 10 times higher than in our human study, was found to be associated with a time-dependent mode with increased secretion of hepatic PLA2 which was exacerbated in the absence of hepatic COX-2 [37]. Thus, the temporary increase in cis-EpOA and 15(S)-8-iso-PGF2α observed in our study may be due to paracetamol-induced short-term hepatotoxicity in the healthy subjects.

4.5. Effects of Paracetamol on Oxidative Stress

In the human study, paracetamol (3 g) did not increase oxidative stress as assessed by measuring urinary excretion of the oxidative stress biomarker 15( )-8-iso-PGF2α [21, 22]. As discussed above, the sharp and short-lasting increase in the plasma concentration of free 15( )-8-iso-PGF2α is likely to be due to temporary release of PLA2 from the liver and/or due to activation of extracellular PLA2. At this high dose, paracetamol did not increase plasma total hCys which is generally assumed to be associated with oxidative stress. Given the ROS-scavenging phenolic moiety of paracetamol ( -acetyl- -aminophenol), the failure of paracetamol to enhance oxidative stress seems reasonable. The F2-isoprostane 15( )-8-iso-PGF2α is known to be produced from AA by the catalytical action of COX [38]. In contrast to acetylsalicylic acid, indomethacin, and celecoxib [25, 39], our study indicates that paracetamol (3 g) does not inhibit COX-dependent formation of 15( )-8-iso-PGF2α in humans.

5. Conclusion

We investigated in vitro and in vivo effects of paracetamol, an analgesic and antipyretic phenolic drug, on the L-Arg/NO, AA/COX, and CYP biochemical pathways and on oxidative stress. At the high single oral dose of 3 g, paracetamol did not alter oxidative stress in vivo. At suprapharmacological concentrations, paracetamol also did not alter oxidative stress in vitro as revealed by the unchanged nitrite-to-nitrate molar ratios measured in incubation mixtures of recombinant heNOS and in cultures of adult rat hepatocytes that express iNOS. The potent PGI2 inhibition by high-dosed paracetamol in the healthy subjects of the present study suggests that the relatively small increase in blood pressure seen in CAD patients by others [9] is likely to be due to compensatory mechanisms that involve enhanced formation of vasodilators. Potential candidates are NO and epoxyeicosatrienoic acids (EETs). In the circulation, NO can be produced from L-arginine by the catalytic action of eNOS and/or from nitrite/nitrate. EETs are produced from arachidonic acid by the catalytic action of the CYP family [34]. Our results suggest that in healthy subjects NO may compensate the loss of the vasodilatory and antiaggregatory prostacyclin caused by high-dosed paracetamol (Figure 8). Also, paracetamol does not increase oxidative stress even when given at suprapharmacological doses. We assume that in healthy humans the paracetamol-induced shift of the PGI2/TxA2 balance is counteracted by concomitant increase in circulating NO production. The underlying mechanisms remain elusive. Possible contributing mechanisms may include elevation of NO in the endothelium and conversion of nitrate to nitrite and its consecutive reduction to NO. In patients suffering from cardiovascular diseases, that is, with dysfunctional endothelium, paracetamol only partially counteracts its unfavorable vasodilatory/vasoconstrictory effect via NO.


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