How can the aerobe Pseudomonas aeruginosa grow in the absence of oxygen?

Answer 1

The piece of research above (which is unlikely to have even been read by the poster) doesn't answer the question.

If Pseudomonas aeruginosa grows both in the prescence and abscence of oxygen, then it is a facultative anaerobe, meaning it can do both. However, it is likely that it didn't grow so well without oxygen.

If all literature says its an obligate areobe, and you got it to grow in anaerobic conditions then you should:
Re-do the experiment because you are likely to have done something wrong

and if you get the same result:
Get it published so everyone knows!

Answer 2

Pseudomonas Aeruginosa Anaerobic

Answer 3

Stratified patterns of protein synthesis and growth were demonstrated in Pseudomonas aeruginosa biofilms. Spatial patterns of protein synthetic activity inside biofilms were characterized by the use of two green fluorescent protein (GFP) reporter gene constructs. One construct carried an isopropyl-β-d-thiogalactopyranoside (IPTG)-inducible gfpmut2 gene encoding a stable GFP. The second construct carried a GFP derivative, gfp-AGA, encoding an unstable GFP under the control of the growth-rate-dependent rrnBp1 promoter. Both GFP reporters indicated that active protein synthesis was restricted to a narrow band in the part of the biofilm adjacent to the source of oxygen. The zone of active GFP expression was approximately 60 μm wide in colony biofilms and 30 μm wide in flow cell biofilms. The region of the biofilm in which cells were capable of elongation was mapped by treating colony biofilms with carbenicillin, which blocks cell division, and then measuring individual cell lengths by transmission electron microscopy. Cell elongation was localized at the air interface of the biofilm. The heterogeneous anabolic patterns measured inside these biofilms were likely a result of oxygen limitation in the biofilm. Oxygen microelectrode measurements showed that oxygen only penetrated approximately 50 μm into the biofilm. P. aeruginosa was incapable of anaerobic growth in the medium used for this investigation. These results show that while mature P. aeruginosa biofilms contain active, growing cells, they can also harbor large numbers of cells that are inactive and not growing.

Biofilms are communities of microorganisms that are associated with surfaces. Cell clusters in biofilms are known to be characterized by gradients in the concentrations of oxygen, nutrients, and metabolic waste products. It is widely recognized that such chemical heterogeneity in microbial biofilms can lead to microorganisms in the biofilm exhibiting different rates of growth and metabolic activity. Even pure cultures of microorganisms growing in biofilms experience these gradients and may exist in a range of metabolic states. The variety of growth states that can be represented in a biofilm, even for a single species, surely contributes to the special ecology and antimicrobial tolerance manifested by biofilms. Given the fundamental significance of growth status, it is surprising that there have been few investigations in which the growth patterns in biofilms have been visualized.

Several studies have investigated spatial patterns of cellular activity inside biofilms, using approaches such as staining with nucleic acid dyes that differentially indicate DNA and RNA (17), hybridization to 16S rRNA with fluorescently tagged oligonucleotide probes (9), the induction of alkaline phosphatase followed by staining with a fluorogenic phosphatase substrate (8, 19), and green fluorescent protein (GFP) expression from a growth-rate-dependent promoter (11). These previous investigations have revealed gradients in metabolic activity in biofilms.

The purpose of the work reported here was to evaluate and apply fluorescent protein-based approaches for mapping spatial patterns of protein synthetic activity in Pseudomonas aeruginosa biofilms. We tested three different fluorescent protein-based approaches for visualizing patterns of activity in P. aeruginosa biofilms. The first used the gfpmut2 gene controlled by the isopropyl-β-d-thiogalactopyranoside (IPTG)-inducible Ptrc promoter. This led to the production of a stable GFP when the gene was induced. The second used a protease-sensitive GFP encoded by a gfp derivative under the control of a growth-rate-dependent promoter, rrnBp1. The third used a fluorescent timer protein that turns from green to red as the protein matures. The activities reflected by these fluorescent protein constructs were determined by confocal scanning laser microscopy of two different biofilm systems.

Bacterial strains and media. P. aeruginosa PAO1 and its derivatives were used for these studies. P. aeruginosa strain AH298 contains a chromosomally encoded gene for GFP under the control of the rrnBp1 promoter. This strain was constructed as described elsewhere (10, 11), except that the kanamycin resistance marker was replaced with tellurite resistance (150 μg/ml) for the selection of transconjugants of P. aeruginosa. Because expression from the rrnBp1 promoter is growth rate dependent (11) and because the expressed GFP is an unstable variant (gfp-AGA), this reporter has the effect of yielding bright green fluorescence in rapidly growing cells and dim or no fluorescence in slowly growing bacteria or bacteria that are not growing (11). Plasmid pAB1 carries gfpmut2 under the control of the IPTG-inducible Ptrc promoter (16). This GFP has enhanced fluorescence, as described by Cormack et al. (4). P. aeruginosa expressing gfpmut2 has been shown to be brightly fluorescent (11). Plasmid pMF335 contains the gene for the timer fluorescent protein (14), a derivative of the Discosoma red fluorescent protein that changes from green to red fluorescence as the protein matures. This plasmid was constructed by ligating the pRO1614 origin of replication from plasmid pMF17 (6) to the unique EcoRI site of pTIMER (BD Biosciences Clontech, Palo Alto, Calif.). The pMF335 construct allowed the stable replication of pTIMER in P. aeruginosa and the expression of the fluorescent timer protein under the control of the Plac promoter. Plasmids pMF335 and pAB1 were introduced into P. aeruginosa by triparental mating with the conjugation helper plasmid pRK2013 (5). Transconjugants were selected on Pseudomonas isolation agar (Difco) containing carbenicillin (300 μg/ml). Bacteria were grown in tryptic soy broth (TSB) or on tryptic soy agar (TSA) plates at 37°C. One-tenth strength TSB was used for flowthrough biofilm experiments.

GFP induction in planktonic cells. Overnight planktonic cultures of P. aeruginosa PAO1(pAB1) were grown with shaking in TSB supplemented with 150 μg of carbenicillin/ml. These cultures were diluted to an optical density (at 600 nm, with a 1-cm path length) of 0.040 in 150 ml of fresh TSB. The cultures were incubated at 37°C and sampled every 20 min throughout the duration of the experiment. At each time point, 3.6 ml of culture was sampled from the flask and pipetted in 200-μl aliquots into a black-sided clear-bottomed 96-well plate (Costar; Corning Incorporated, Corning, N.Y.). Fluorescence intensities were measured with a Bio-Tek FL600 microplate fluorescence reader, with excitation at 485 nm and emission at 530 nm. The fluorescence intensity values were averaged for 16 samples for each time point. After 2 h, the culture was divided into two equal volumes. The bacteria in one flask were induced with 1 mM IPTG, while those in the other flask served as a control without induction.

For tests of GFP induction in an anaerobic environment, nitrogen gas was bubbled through TSB in two anaerobic vials. The vials were sealed and autoclaved. Bacteria from aerated overnight cultures of P. aeruginosa PAO1(pAB1) in TSB with 150 μg of carbenicillin/ml were injected into the anaerobic TSB to achieve a final optical density of 0.040. The beginning culture volume in each vial was approximately 80 ml. The vials were sampled every 30 min for 2 h with sterile needles and syringes that were purged with nitrogen. GFP fluorescence intensities were determined for these samples as described above. After 2 h, IPTG was added to one vial to a final concentration of 1 mM. The same amount of sterile buffer was added to the control. Sampling continued as described above every 30 min for 4 h.

Colony biofilm preparation. Colony biofilms were grown on polycarbonate membranes resting on agar plates (1, 16). Planktonic cultures of P. aeruginosa were grown overnight with shaking in TSB and diluted to an optical density (at 600 nm, with a 1-cm path length) of 0.050 in TSB. Carbenicillin (150 μg/ml) was added to the broth used to grow the inoculum for the preparation of PAO1 with fluorescent timer protein, PAO1(pMF335), or PAO1(pAB1). The antibiotic was not included in the agar medium used to grow colony biofilms. One 5-μl drop of diluted planktonic culture was used to inoculate individual sterile, black, polycarbonate membrane filters (25-mm diameter, 0.2-μm pore size; Poretics Corp., Livermore, Calif.) resting on TSA. The membranes were sterilized by UV exposure (15 min per side) prior to inoculation. The plates were inverted and incubated at 37°C for 48 h, and the membrane-supported biofilms were transferred to fresh culture medium once after 24 h. For the strain with the IPTG-inducible GFP, the biofilms were transferred to agar medium containing 1 mM IPTG for an additional 4 h. To confirm the presence of bacteria in the uninduced region of the PAO1(pAB1) colony biofilms, we used a rhodamine counterstain. Rhodamine B (5 μg/ml) was added to the TSA plates used for the final 4 h of incubation to counterstain bacteria that did not express GFP.

For anaerobic control experiments, TSA plates were incubated in anaerobic bags with sealing bars (BBL GasPak system; Becton Dickinson Microbiology, Franklin Lakes, N.J.) for 24 h prior to membrane inoculation. Colony biofilms were prepared as described above except that immediately after membrane inoculation, the plates were again placed inside anaerobic bags for incubation at 37°C. At the time of sampling, only the plate being sampled was removed from its anaerobic bag.

Freeze sectioning and microscopy. Colony biofilms were embedded in a histological cryoembedding medium (Tissue-TEK O.C.T. compound; VWR Scientific Products, Willard, Ohio), and dry ice was used to solidify the embedding medium. Membrane colony biofilms were placed on a flat piece of flexible metal, and the colonies were covered with embedding medium. The metal was placed on dry ice and the embedding medium was allowed to freeze. Membranes, colonies, and the embedding medium were detached from the metal sheet, and the edges of the membranes were trimmed. The colonies were then placed directly on dry ice, with the embedded side down and the exposed membrane side up. Embedding medium was used to cover the exposed membranes, which were then frozen.

Embedded colonies were sectioned into 5-μm-thick cross sections by use of a Leica CM1800 refrigerated microtome. The sections were placed on glass slides and then examined by confocal scanning laser microscopy.

Capillary biofilm system. Biofilms were grown in glass capillary tubes under continuous flow conditions (Fig. 1). The glass tubes had square cross sections, allowing direct microscopic observation of the biofilms growing on the inside of the tubes through the flat tube walls. The capillaries were mounted in a flow cell holder to reduce breakage. The capillaries had a nominal inside dimension of 900 μm and a wall thickness of 170 ± 10 μm (Friedrich & Dimmock, Millville, N.J.). The flow cell apparatus consisted of a vented medium feed carboy (4-liter capacity), a flow break, a filtered air entry, a peristaltic pump, the capillary and flow cell holder, an inoculation port, and a waste carboy. These components were connected by silicone rubber tubing. We have found that the development of biofilm cell clusters of P. aeruginosa is enhanced by pumping air through the capillary along with the liquid medium. Therefore, the system also contained a T connector just upstream of the glass capillary to allow mixing of the air and medium flows. Medium and system components were sterilized separately by autoclaving and then connected after cooling in a biological hood.

The capillary flow system was inoculated with 2 ml of an overnight culture of P. aeruginosa with an optical density at 600 nm of 0.001 to 0.005. For inoculation, the flow was stopped and the tubing was clamped downstream of the inoculation port. The inoculum was injected via the port to fill the glass capillary. The tubing upstream of the glass tube was clamped, and the system was allowed to stand without flow for 24 h. After this time, the flow of medium (1/10-strength TSB) was initiated at a flow rate of 20 ml h−1. Air was pumped through the capillary at the same flow rate as the medium by use of a parallel tube in the same peristaltic pump. This resulted in slug flow of the medium and air bubbles through the capillary tube. Biofilms were counterstained by injecting a solution of rhodamine B at 50 mg liter−1 into the capillary. Biofilms were observed by scanning confocal laser microscopy after 24 h of continuous flow at 37°C.

Microscopy and image analysis. Confocal scanning laser microscopy was performed with a Leica TCS NT confocal scanning laser microscope, with excitation at 488 and 568 nm and with emission collected at 500 to 530 nm (green channel) and 585 to 615 nm (red channel). Microscope images were analyzed by use of the line-scan function of MetaMorph image analysis software (Universal Imaging Co., Downingtown, Pa.). For colony biofilm experiments, the relative GFP intensities were measured for at least three independently grown biofilms and for three independent transects across each biofilm. Depth profiles of GFP intensity were measured perpendicular to the membrane supporting the colony biofilm. The interface between the biofilm and the membrane was set at zero on this spatial axis. Intensity profiles were used to measure the biofilm thickness and the dimensions of the green fluorescent band of active growth produced by GFP from strains PAO1(pAB1) and AH298. The biofilm thickness was measured as the distance from the membrane to the biofilm-air interface.

Oxygen penetration. Oxygen concentration profiles in colony biofilms were measured with a dissolved oxygen microelectrode. The oxygen microelectrode was based on the principle of the common amperometric Clark oxygen electrode. It consisted of an outer casing sealed at the sensor tip with an oxygen-permeable silicone membrane. The casing was fabricated from a Pasteur pipette that was tapered down to an active sensor tip of 15 μm. A buffered electrolyte solution consisting of 0.3 M K2CO3, 0.2 M KHCO3, and 1 M KCl filled the internal cavity. Three electrodes occupied the internal cavity as well: they included a gold-tipped, glass-encased platinum cathode at which oxygen diffusing in through the silicone membrane was reduced; a silver-silver chloride counterelectrode, which served as the current return; and a guard electrode, which reduced the amount of unwanted oxygen entering from the back of the electrode. A potential of −0.8 volts of direct current was applied between the cathode and the counterelectrode. The current from the cathode, which is proportional to the concentration of oxygen in the bulk external solution, was measured with a picoammeter in the range of 0 to 3 nA. The same potential was applied between the guard electrode and the counterelectrode to reduce the background signal while measuring low concentrations of oxygen. The electrode was calibrated in air, and a zero level was obtained by placing the electrode tip in a 0.5% agar gel containing a suspension (0.2%) of ferrous sulfide (3). Oxygen concentration profiles were obtained by lowering the oxygen microelectrode into the biofilm by use of a computer-controlled stepping motor (16).

Transmission electron microscopy. Colony biofilms were grown for 48 h on TSA and then exposed to 150 μg of carbenicillin/ml for 12 h by the transfer of biofilms to TSA plates supplemented with the antibiotic. An untreated control was grown for 60 h on TSA. These specimens were fixed overnight by moving the colony biofilm to a filter pad soaked in 5% glutaraldehyde. The biofilms were postfixed in 2% osmium tetroxide, dehydrated in an ethanol series, and treated with propylene oxide. Colonies were embedded in Spurr's epoxy resin. Ultrathin sections were cut, placed on copper grids, and stained with uranyl acetate and Reynold's lead citrate. The sections were examined on an LEO 912 AB transmission electron microscope. Cell lengths were measured and displayed as a function of their relative positions in the biofilm. Cell lengths, measured along the major axis of the cell, were measured manually from enlarged prints.

Stratified pattern of GFP fluorescence in P. aeruginosa colony biofilms. P. aeruginosa formed colony biofilms that were approximately 150 to 300 μm thick after 48 h of development (Table 1). Cell numbers in colony biofilms were similar for strains PAO1, PAO1(pAB1), and AH298, with a mean viable areal cell density of approximately 2 × 109 CFU cm−2. Strain PAO1(pAB1) carries a plasmid containing a gfpmut2 gene, which encodes a stable GFP that is induced in the presence of IPTG. The addition of IPTG to an aerobic exponential-phase planktonic culture resulted in the development of green fluorescence in the culture, as determined by fluorimetry (Fig. 2A). No increase in fluorescence was observed in the absence of IPTG or in cells cultivated under anaerobic conditions (Fig. 2B). Optical density measurements conducted in parallel showed that P. aeruginosa grew under aerobic conditions, but not under anaerobic conditions, in TSB (Fig. 3). When P. aeruginosa PAO1(pAB1) was cultivated in colony biofilms under uninduced conditions, no green fluorescence was observed. Figure 4A shows an uninduced PAO1(pAB1) colony biofilm counterstained with rhodamine B. In contrast, bright green fluorescence located all along the air interface of the colony biofilm was visible in PAO1(pAB1) colony biofilms that were exposed to IPTG for 4 h (Fig. 4B).

Similar results were observed with P. aeruginosa AH298, which carries a gfp derivative encoding an unstable GFP under control of the growth-rate-dependent rrnBp1 promoter. For this strain, rapidly growing cells are bright green, while slowly growing cells are dim or dark. In colony biofilms formed by strain AH298, green fluorescence was localized in a band adjacent to the air interface of the colony (Fig. 4C). No such distinct band could be discerned in biofilms formed by wild-type PAO1 (not shown).

We also attempted to visualize spatial patterns of activity by using a fluorescent timer protein construct. This strain carries a plasmid coding for a fluorescent protein that has been shown to change from green to red fluorescence in an oxygen-dependent maturation process as the protein ages. One possibility was that this construct would identify areas of new growth in green and areas of aged cells in red. Both green and red fluorescence was observed in colony biofilms, but the two colors were distributed uniformly throughout the depth of the biofilm (not shown).

The bands of GFP expression for strain PAO1(pAB1) and AH298 occupied about one-third to one-fourth of the colony thickness. The dimension of the fluorescent region averaged 60 ± 12 μm for PAO1(pAB1) and 63 ± 16 μm for AH298 (Table 1). The zone of active protein synthesis revealed by these constructs coincided with the aerobic layer of the biofilm, as determined by the use of oxygen microelectrodes (Fig. 5). Oxygen penetrated about 50 μm into the colony biofilms formed by any of the P. aeruginosa strains (Table 1). The measured depth of oxygen penetration was not statistically significantly different for PAO1, AH298, or PAO1(pMF335) colony biofilms (P > 0.35 by a two-tailed t test).

P. aeruginosa does not grow on TSA under anaerobic conditions. GFP requires oxygen for the activation of fluorescence (15). In a biofilm that may contain anoxic zones, a lack of fluorescence could be due to an absence of protein synthetic activity, but it is also possible that the protein was made and simply did not mature to a fluorescent state because of the local absence of oxygen. To help discern these possibilities, we tested the growth of P. aeruginosa under anaerobic conditions without the addition of an alternative electron acceptor. No growth was detected in anaerobically incubated batch planktonic cultures (Fig. 3A). When P. aeruginosa was grown aerobically on TSA, the viable cell numbers in a colony biofilm increased during 48 h of development, from 8 × 104 to 5 × 109 CFU. When the strains were incubated anaerobically on TSA, the viable cell numbers increased very little, from 8 × 104 CFU at the time of inoculation to 1.3 × 105 CFU. No growth was visible when P. aeruginosa was streaked onto a TSA plate and incubated anaerobically, even after several days. These observations confirm that P. aeruginosa is not capable of anaerobic growth on the medium that we used.

Stratified pattern of cell elongation in antibiotic-treated colony biofilms. When P. aeruginosa is exposed to the antibiotic carbenicillin, cell division is blocked and growing cells form filaments. We used this behavior as another means to localize regions of growth in colony biofilms. Transmission electron microscopy of colony biofilms left untreated for 48 h showed short rods, which are characteristic of P. aeruginosa (Fig. 6, top row). The untreated biofilms also harbored significant numbers of lysed cells. Lysed cells predominated in the colony interior, with fewer lysed cells near the air interface. When 48-h colony biofilms were exposed to 150 μg of carbenicillin ml−1 for an additional 12 h, filamentous cells were evident, but only near the colony-air interface (Fig. 6, bottom left panel). Cell elongation was not observed for cells in the interior of the carbenicillin-treated biofilms or near the membrane (Fig. 6, bottom right panel). Filamentation was observed on the opposite interface of the colony from where the antibiotic was delivered, showing that carbenicillin penetrated the biofilm but only affected cells at the air interface. The distribution of elongated cells in the antibiotic-treated specimen and in the untreated control is shown in Fig. 7. Since cell elongation depends on growth, this result demonstrates that growth occurs in a layer of the biofilm near the air interface and suggests that there is no growth in strata of the biofilm that are more distant from the oxygen source.

Stratified pattern of GFP fluorescence in P. aeruginosa capillary tube biofilms. To determine whether the inducible GFP reporter described above could be applied in a noninvasive manner, we induced P. aeruginosa PAO1(pAB1) biofilms developed in glass capillary tubes with IPTG for 4 h and then examined them for fluorescence by using confocal scanning laser microscopy. These experiments differed from those with colony biofilms in that there was a continuous flow of medium and the biofilms did not have to be frozen and sectioned prior to examination. The same medium and temperature were used to grow biofilms in capillary tubes as were used for colony biofilms.

After 24 h of growth, large clusters of bacterial microcolonies were evident on the walls of the capillaries (Fig. 8A). The expression of GFP was limited to the surfaces of these clusters and was primarily evident in the vicinity of interstices (Fig. 8B). The dimension of the zone of GFP expression in this system was approximately 25 to 30 μm.

Attempts to map spatial patterns of growth in the presence of nitrate were not fruitful. We attempted to measure the patterns of GFP expression and cell elongation in P. aeruginosa colony biofilms growing on TSA supplemented with nitrate. One would anticipate that the addition of nitrate would lead to growth in anaerobic layers of the biofilm that were incapable of growth in the absence of nitrate in earlier experiments. But because GFP activation is oxygen dependent, we could also predict that this growth would not be visible, i.e., anaerobic synthesis of GFP would occur, but the GFP would not become fluorescent. When PAO1(pAB1) was grown on TSA supplemented with 1% potassium nitrate and induced with IPTG, GFP induction along the air interface of the colony was observed as usual. No additional expression of GFP was visible elsewhere in the colony. There was no indication of GFP expression along the agar interface of the colony, which was adjacent to the source of nitrate. We also attempted to localize growth in colony biofilms growing on TSA supplemented with nitrate by examining specimens of cell elongation after exposure to carbenicillin. No filamentous cells were seen in such samples, not even at the air interface of the biofilm. Since we know that viable cell numbers increase in colony biofilms grown on media containing nitrate, we concluded that nitrate interferes with the filamentation that is normally induced by carbenicillin treatment. In summary, attempts to map spatial patterns of growth in biofilms growing on a medium containing nitrate were not fruitful. Neither the inducible GFP nor the cell elongation techniques were able to provide information about the pattern of growth in P. aeruginosa colony biofilms grown in the presence of nitrate that could be unambiguously interpreted.