Wellman Laboratories of Photomedicine, Massachusetts General Hospital and Department of Dermatology, Harvard Medical School, Boston, MA 02114-2698, USA
Received 5 November 2001; returned 24 January 2002; revised 25 February 2002; accepted 8 March 2002.
![]() |
Abstract |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Methods: Conjugates were prepared by attaching precisely one chlorine6 molecule to the -amino group of poly-(
-benzyloxycarbonyl)lysines of average length eight and 37 lysine residues, followed by deprotection of the
-amino groups, and were characterized by iso-electric focusing. The uptake of these conjugates and free chlorine6 by Gram-positive Staphylococcus aureus (ATCC 27659) and Gram-negative Escherichia coli (ATCC 29181) after washing was measured as a function of photosensitizer concentration (04 µM chlorine6 equivalent) and incubation time. After incubation the bacteria were exposed to low fluences (1040 J/cm2) of 660 nm light delivered from a diode laser, and viability was assessed after serial dilutions by a colony-forming assay.
Results: S. aureus and E. coli took up comparable amounts of the two conjugates, but free chlorine6 was only taken up by S. aureus. After illumination S. aureus was killed in a fluence-dependent fashion when loaded with the 8-lysine conjugate and free chlorine6 but somewhat less so with the 37-lysine conjugate. In contrast, PDI of E. coli was only effective with the 37-lysine conjugate at concentrations up to 4 µM. PDI using the 8-lysine conjugate and free chlorine6 on E. coli was observed at a concentration of 100 µM. Transmission electron micrographs showed internal electron-lucent areas consistent with chromosomal damage.
Conclusion: These results can be explained by the necessity of a large polycation to penetrate the impermeable outer membrane of Gram-negative E. coli, while Gram-positive S. aureus is more easily penetrated by small molecules. However, because S. aureus is more sensitive overall than E. coli the 37-lysine conjugate can effectively kill both bacteria.
Keywords: photodynamic therapy, polylysine, Escherichia coli, Staphylococcus aureus, electron microscopy
![]() |
Introduction |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Many naturally occurring antibacterial peptides have been discovered that all have in common inter alia a pronounced polycationic charge.19 This is thought to be the initial factor that allows them to bind to negatively charged bacteria and subsequently disturb the outer-membrane permeability barrier.20 Antibacterial polycations include polymyxins,21 protamine,22 insect cecropins,23 reptilian magainins,24 various cationic leucocyte peptides (defensins,25 bactenecins,26 bactericidal/permeability-increasing protein27), polymers of basic amino acids28 and polyethyleneimine.29 However, the cationic character is not the sole determinant required for the permeabilizing activity, and therefore some of the agents are much more effective permeabilizers than others.
We showed previously30 that a polycationic covalent conjugate between the PS chlorine6 (ce6) and poly-L-lysine (pL) (average molecular weight 2 kDa, 20 lysine residues) effectively delivered PS to both Gram-positive and Gram-negative oral bacteria after a short incubation, and enabled their photodestruction after illumination with red light. The activity of the cationic pL-ce6 was many times higher than the neutral acetylated pL-ce6-ac and anionic succinylated pL-ce6-succ conjugates against both Gram-positive and Gram-negative bacteria. With the relatively short incubation times used in this study, mammalian epithelial cells were essentially unharmed by the treatment. Other workers subsequently used conjugates between porphycenes and pL to carry out PDI of both Gram-positive and Gram-negative bacterial species. In order for bacterial PDI to have any clinical application it is necessary to demonstrate that a single conjugate can kill both classes of bacteria, so that the method can be used without prior identification of the infectious agent.
This report tests the hypothesis that a certain length of polycationic chain is necessary to allow the PS to gain entry through the outer membrane of Gram-negative bacteria, while this large polycationic carrier may be unnecessary or even detrimental with Gram-positive bacteria. In order to produce conjugates with a more defined structure, a synthetic strategy was devised that enabled precisely one ce6 molecule to be attached to the pL chain.
![]() |
Materials and methods |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
All reactions were carried out in the dark at room temperature. Forty-seven milligrams (4.8 µmol) of poly--(benzyloxycarbonyl)-DL-lysine [average molecular weight = 9700 (range = 500020 000), mean degree of polymerization = 37; Sigma Chemical Co., St Louis, MO, USA] were dissolved in 1 mL of dry dimethylsulphoxide (DMSO), to which 6.7 mg (11.2 µmol) of ce6 (Porphyrin Products, Logan, UT, USA) and 30 mg (157 µmol) of 1-ethyl-3-(3-dimethylaminopropyl)-carbodiimide hydrochloride (Sigma) were added. Triethylamine (20 µL; Sigma) was then added and the mixture was stirred for 24 h. Likewise, 40 mg (20 µmol) of poly-
-(benzyloxycarbonyl)-L-lysine [average molecular weight = 2000 (range = 10004000), mean degree of polymerization = 8] were reacted with 28 mg (47 µmol) of ce6 and 125 mg (654 µmol) of 1-ethyl-3-(3-dimethylaminopropyl)carbodiimide hydrochloride (EDC). The reaction was checked by thin layer chromatography (Polygram SIL G/UV254 plates; Macherey Nagel, Duren, Germany) in methylene chloride/methanol/ammonium hydroxide (8:2:0.5). After 24 h when the reaction was complete, methanol (1 mL) and water (1 mL) were added and the mixture was evaporated to dryness under vacuum. Trifluoroacetic acid (1 mL) was then added to the dry mixture and this solution was stirred for 2 h. Sodium acetate buffer (10 mM, pH = 5.5, 1 mL) and methanol (1 mL) were then added and the mixture was again evaporated to dryness under vacuum. The residue was dissolved in sodium acetate buffer (10 mM, pH = 5.5, 1 mL) and applied to a column of Sephadex G25 (60 x 1 cm) and eluted with sodium acetate buffer (10 mM, pH = 5.5) at a flow rate of 3.3 mL/h. Three millilitre fractions were collected and fractions 1421 were combined and evaporated to give the product pL-ce6. Concentrations of free ce6 and conjugates were measured routinely by absorption spectroscopy taking the extinction coefficient
400nm = 150 000. Isoelectric focusing (IEF) was carried out in agarose gels containing wide range ampholines (pI 312; Sigma) that were subjected to pre-focusing at 800 V for 30 min. Samples were then loaded followed by a desalting period of 150 V for 30 min and a focusing period at 1400 V for 60 min. ce6 was localized on the gel by fluorescence imaging using an excitation 400440 nm bandpass filter, and an emission 580 nm longpass filter (ChemiImager 4000; Alpha Innotech Corp, San Leandro, CA, USA). pL was localized by Coomassie blue staining.
Bacteria
The bacteria used in this study were Staphylococcus aureus (ATCC 27659, penicillinase-positive, resistant to tetracycline, novobiocin, streptomycin and macrolide antibiotics) and Escherichia coli K12 (ATCC 29181, resistant to trimethoprim and streptomycin). The bacteria were grown in brain-heart infusion broth in an orbital shaker at 37°C for 18 h. An aliquot of this suspension was then added to nutrient broth and grown to mid-log phase (OD600 = 0.6, 108 cells/mL).
Uptake studies
Bacteria were used at a density of 109 cells/mL (prepared by concentrating a suspension of 108 cells/mL) in order to have sufficient material to measure the extracted ce6. Bacteria were incubated in the dark at room temperature for 30 min with various concentrations of conjugates or free ce6 measured as µM ce6 equivalent (final concentration in nutrient broth). Experiments were carried out in triplicate. The cell suspensions were centrifuged (9000g, 1 min), the PS solution was aspirated and bacteria were washed twice by resuspending the cell pellet in 1 mL of sterile phosphate-buffered saline (PBS) and centrifuged as above. Finally, the cell pellet was dissolved by digesting it in 1.5 mL of 0.1 M NaOH/1% SDS for at least 24 h to give a homogeneous solution. The fluorescence of the cell extract was measured on a spectrofluorimeter (model FluoroMax; SPEX Industries, Edison, NJ, USA). The excitation wavelength was 400 nm and the emission spectra of the cell suspensions were recorded from 580 to 700 nm. If necessary, the solution was diluted with 0.1 M NaOH/1% SDS to reach a concentration of ce6 where the fluorescence response was linear. Separate fluorescence calibration curves were constructed with known amounts of both conjugates and free ce6 dissolved in 0.1 M NaOH/1% SDS. The protein content of the entire cell extract was then determined by a modified Lowry method31 using bovine serum albumin dissolved in 0.1 M NaOH/1% SDS to construct calibration curves. Results were expressed as nmol ce6/mg cell protein.
PDI studies
Suspensions of bacteria (108/mL) were incubated in the dark at room temperature for 30 min with 0.54 µM ce6 equivalent of the two conjugates and free ce6 in nutrient broth as described above. Cell suspensions were centrifuged, cells were washed twice with sterile PBS and 1 mL of fresh nutrient broth was added. The bacterial suspensions (1 mL) were then placed in the wells of 24-well plates. The wells were illuminated from below in the dark at room temperature. A 660 nm, 300 mW diode laser (SDL Inc, San Jose, CA, USA) was coupled into a 1 mm optical fibre that delivered light into a lens that formed a uniform circular spot on the base of the 24-well plate 2 cm in diameter. Fluences ranged from 0 to 40 J/cm2 at an irradiance of 50 mW/cm2. The plates were kept covered during the illumination in order to maintain the sterility of the culture. At intervals during the illumination when the requisite fluences had been delivered, aliquots (100 µL) were taken from each well to determine colony-forming units (cfu). Care was taken to ensure that the contents of the wells were mixed thoroughly before sampling, as bacteria can settle at the bottom. The aliquots were serially diluted 10-fold in nutrient broth to give dilutions of 101106 times the original concentrations. Aliquots (10 µL) of each of the dilutions were streaked horizontally on square nutrient agar plates as described by Jett et al.32 Plates were streaked in triplicate and incubated for 24 h at 37°C in the dark. In general, three dilutions could be counted on each plate. Controls were bacteria untreated with PS or light but kept in 24-well plates at room temperature covered with aluminium foil for the duration of the illumination, and bacteria exposed to light in the absence of PS.
Incubation time
A preliminary experiment was carried out in order to determine the effect of the incubation time on the efficiency of the PDI of bacteria. S. aureus and E. coli (108 cells/mL) were incubated with the 37-lysine conjugate (1 µM ce6 equivalent in nutrient broth) for times ranging from 1 to 30 min. Two aliquots were removed at each time point: one for the determination of PDT-mediated killing, and the other for the measurement of dark toxicity, both measured by colony-forming assay as described above.
Transmission electron microscopy
Control and conjugate-treated bacteria (both dark- and light-treated) were suspended in PBS. PDI conditions were 4 µM ce6 equivalent and 20 J/cm2. The bacteria were pelleted, PBS decanted and fixed in 4% glutaraldehyde in 0.1 M cacodylate buffer (pH 7.4) for 1 h at 4°C. The bacterial pellets were transferred to 1.5 mL microcentrifuge tubes and centrifuged for 5 min at 12 000g. The glutaraldehyde was decanted, the pellets rinsed twice in 0.1 M cacodylate buffer and the bacteria spun down again. After decanting the buffer, the bacteria were resuspended in 2% molten Bacto agar (Difco) and centrifuged immediately. The agar pellets containing bacteria were cut into 3 mm3 pieces and processed for routine electron microscopy. The bacterial pellets were co-fixed in 2% OsO4, dehydrated in alcohols and embedded in Epon 812. Thin sections were cut on an ultramicrotome, stained with uranyl acetate and lead citrate, and examined on a transmission electron microscope.
![]() |
Results |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Because Sigma offered two lengths of poly-(-benzyloxy-carbonyl)-L-lysine with average chain lengths of eight and 37 lysines, it was decided to compare the effect of chain length on the photosensitizing effects of pL-ce6. We decided to use EDC as the coupling agent with free ce6 in DMSO in an attempt to avoid problems of aggregate formation. The results of the synthesis were favourable in that conjugates were obtained that could be purified easily by column chromatography and more importantly did not show any time-dependent aggregation behaviour.
The conjugates were characterized by IEF, taking advantage of their charged nature to effect separation, and the ability to localize ce6 independently by fluorescence imaging, and pL by Coomassie blue staining on the gel. Fluorescence and Coomassie images of the two conjugates and free ce6 are shown in Figure 1. The 37-lysine conjugate produced a streak that encompasses a higher range of pI values than the 8-lysine conjugate, while free ce6 had a low pI value of 34. The streaking of the 37-lysine conjugate is presumably due to the greater degree of polydispersity inherent in larger pL chains compared with small ones. In order to demonstrate that ce6 is attached covalently to the pL in the conjugates, we also loaded a mixture of pL (8 lysines) and free ce6 in lane 1. It can be seen that the ce6 fluorescence was well separated from the pL chain.
|
In order to decide on an incubation time for subsequent experiments we investigated the effect of increasing the incubation time of the two bacteria with the 37-lysine conjugate at 2 µM ce6 equivalent in nutrient broth. On completion of incubations, the bacteria were either illuminated or kept in the dark, and the numbers of cfu were determined. Figure 2 shows that the survival fraction after illumination decreased fairly sharply with increasing incubation time for both species. S. aureus was significantly more sensitive than E. coli both in the light and the dark. Considering these data we decided to use an incubation time of 30 min for subsequent experiments rather than the time of 1 min used in our previous study.
|
The uptake of ce6 from both the conjugates and unconjugated ce6 with increasing PS concentrations after 30 min incubation in nutrient broth is shown in Figure 3a (S. aureus) and Figure 3b (E. coli). The uptake of ce6 from the 8-lysine and 37-lysine conjugates was remarkably similar for both bacteria, and the uptake from the 8-lysine conjugate was significantly higher in both cases than the uptake from the 37-lysine conjugate. The most striking difference between the two species of bacteria was in the uptake of free ce6, where the Gram-positive S. aureus took up more than 10 times as much as the Gram-negative E. coli. There appears to be a tendency for the uptake of all three species by E. coli to saturate with increasing concentration in the medium, as compared with the uptake by S. aureus.
|
PDI of the two bacterial species was then carried out after 30 min incubation in nutrient broth with the three PSs at concentrations of 1, 2 and 4 µM. The illumination was also carried out in nutrient broth. The data are shown in Figure 4 (af). The survival fraction at 0 J/cm2 represents the dark toxicity of the conjugate or PS. Under these conditions the limit of detection of the colony-forming assay was six logs of killing. S. aureus showed a clear PS dose- and light dose-dependent loss of cfu for all three PSs. The highest PS concentration of 4 µM in conjunction with 40 J/cm2 of 660 nm light produced a near six log reduction of cfu in all cases. The lower concentrations of PS (1 and 2 µM) were less effective in the case of the 37-lysine conjugate compared with the killing observed with 1 and 2 µM concentrations of the 8-lysine conjugate and free ce6. The effect of increasing the concentration of PS was less pronounced in the case of free ce6 (Figure 4c) than the two conjugates (Figure 4a and b). There was a small amount of dark toxicity associated with the higher concentrations of the conjugates but none with free ce6. The results were completely different for E. coli. The only phototoxicity observed was with the 37-lysine conjugate (Figure 4d), which showed a PS dose- and light dose-dependent loss of cfu up to three logs. A dose-dependent dark toxicity was observed with survival fractions of 0.43, 0.2 and 0.04 for 1, 2 and 4 µM concentrations. Only the 4 µM concentrations of the 8-lysine conjugate and free ce6 are shown in Figure 4 (e and f) as the lower concentrations had no effect on bacterial survival in either the light or dark.
|
As mentioned earlier, there was no apparent phototoxicity of either pL-ce6 or free ce6 with E. coli, despite the fact that E. coli took up more pL-ce6 8 than pL-ce6 37. The absence of phototoxicity by free ce6 may be explained by the low uptake (less than one-tenth of the value from both the pL-ce6 conjugates). It was, therefore, of interest to increase the concentrations of pL-ce6 8 and free ce6 substantially with E. coli to try to resolve these anomalies. Figure 5a shows the uptake of ce6 by the cells after incubation with concentrations up to 100 µM of both pL-ce6 8 and free ce6 and Figure 5 (b and c) shows the resultant phototoxicity. Both pL-ce6 8 and free ce6 were taken up in a linear fashion with increased concentration, and the ratio of ce6 in cells incubated with both species was similar at 100 µM to that pertaining at 4 µM. When the phototoxicity was studied, it could be seen that, although 100 µM pL-ce6 8 killed up to two logs of bacteria, the increase was much more pronounced in the case of free ce6, with 20 µM killing two logs and 100 µM killing four logs.
|
By defining the phototoxicity as the inverse of the survival fraction and then dividing this by the uptake of ce6 in nmol/mg cell protein and plotting against light fluence, it is possible to depict graphically and compare the efficiency of the different species for PDI of the bacteria in terms of how much killing is accomplished by each molecule of ce6 taken up by the cells. Figure 6b shows that the relative phototoxicity of the 37-lysine conjugate and free ce6 both incubated at 4 µM was identical for S. aureus while that of the 8-lysine conjugate at 4 µM was c. 10-fold higher. For E. coli, since only the 37-lysine conjugate showed phototoxicity at 4 µM, we also plotted the values obtained from free ce6 and pL-ce6 8 at 100 µM. Figure 6a shows that pL-ce6 8 had very low values at both concentrations, while the values for free ce6 showed a remarkable (100 times) rise from 4 to 100 µM. Indeed, the values from free ce6 at 100 µM were somewhat higher than pL-ce6 37 at 4 µM.
|
Figure 7 shows the morphology of S. aureus (Figure 7ae) and E. coli (Figure 7fj) under control conditions, and after treatment with the two conjugates in the dark and exposed to 20 J/cm2 light. S. aureus showed similar internal areas that were electron lucent in cells treated with pL-ce6 37 Lys and light (Figure 7c) and pL-ce6 8 Lys and light (Figure 7e), which were not seen in control cells or cells treated with conjugates in the dark (Figure 7b and d). In contrast, E. coli only showed these internal cavities after treatment with pL-ce6 37 Lys and light (Figure 7h). These electron-lucent areas may be explained by membrane damage leading to chromosomal alteration and DNA condensation. They bear a remarkable resemblance to those transmission electron micrographs presented by Nitzan et al.,14 who used a mixture of polymyxin nonapeptide and deuteroporphyrin to photoinactivate E. coli and Pseudomonas aeruginosa.
|
![]() |
Discussion |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
In our previous publications30,33 we described a synthetic scheme to give pL-ce6 conjugates using dicyclohexylcarbodiimide to prepare an N-hydroxysuccinimide ester from ce6 and its subsequent reaction with pL dissolved in DMSO. However, although these conjugates performed relatively well they suffered from a tendency to aggregate over time (several days) that made purification and storage difficult. In an attempt to avoid this aggregation problem we formed the hypothesis that it might be caused by the structure of the pL-ce6 having ce6 substitution at random points throughout the length of the pL chain. To test this hypothesis we devised a synthetic scheme using a fully -amino-protected pL chain that could therefore only react on the single
-amino group. Not only would this allow substitution of ce6 on the end of the chain instead of throughout its length, but it would also give chains with only one ce6 molecule per pL molecule. This would eliminate the uncertainty caused by random substitution, whereby although one could have an average of one ce6 molecule per chain it is only an average value and some chains could have two while others have none. This method allowed the conjugates to be readily purified using Sephadex chromatography, and they could be characterized using IEF on agarose gels. The 37-lysine conjugate appeared to have a higher pI range than the 8-lysine conjugate, thus providing evidence of its greater polycationic character.
The results of the present study imply that, in the case of polycationic PS conjugates, it is necessary for the PS to gain access through the outer membrane permeability barrier to more sensitive parts of the cell. The efficiency with which this occurs depends on the size of the polycationic chain. All three species (conjugates with eight and 37 lysines and free ce6) efficiently inactivated S. aureus while only the 37-lysine conjugate killed E. coli. The binding of the 8-lysine conjugate to E. coli, however, was higher than that of the 37-lysine conjugate, implying that the 8-lysine conjugate was located at a non-sensitive site. The most obvious explanation of our findings is that the 37-lysine conjugate was able to interact with the outer membrane structure of E. coli, perhaps causing loss of some lipopolysaccharide (LPS) and rendering the remaining LPS more permeable, thus allowing the conjugate to penetrate through the outer membrane to the periplasmic space and to the cytoplasmic membrane beyond. The 8-lysine conjugate, presumably, did not have sufficient polycationic character to accomplish this and, although its overall uptake was higher, the reactive species produced upon illumination were unable to cause lethal damage. In the case of S. aureus it is generally accepted that the peptidoglycan outer layer has much higher permeability34 than the Gram-negative outer envelope, and, although it can carry out molecular sieving for relatively large molecules (up to 50 kDa),35 it presents little barrier to the diffusion of smaller molecules into the periplasmic space. It is possible that the higher uptake and phototoxicity of the 8-lysine conjugate compared with the 37-lysine conjugate could be explained by a molecular-sieving effect that restricts uptake of the larger molecule.
Poly-L-lysines are among the polycations that bind to the anionic sites of LPS. This binding may weaken the intermolecular interactions of the LPS constituents, disorganize the structure and render it permeable to drugs20 by enabling them to cross the outer membrane. Hancock et al.36 coined the term self-promoted uptake to describe the uptake of cationic peptides across outer membranes of Gram-negative bacteria. The first step is the interaction of polycations with divalent cation-binding sites on the cell surface, and since these peptides have much higher affinities for LPS compared with the native divalent cations Ca2+ and Mg2+, they competitively displace these ions and, being so bulky, disrupt the normal barrier property of the outer membrane. The affected membrane is thought to develop transient cracks that permit the passage of a variety of molecules, including hydrophobic compounds and small proteins and/or antimicrobial compounds, and, more importantly, promote the uptake of the perturbing peptide.37
Vaara38 studied the permeabilizing effect on P. aeruginosa of pL chains 3, 4 and 5 lysines in length. Only the 5-lysine chain permeabilized P. aeruginosa but not other Gram-negative species (E. coli or Salmonella typhimurium), and only in low salt and low Mg2+ ion buffers. This was explained by a very weak binding of the 5-lysine chain to the LPS, which could be competitively reversed by Na+ ions as well as Mg2+ ions. The effect of a pL with 20 lysine residues on S. typhimurium with smooth LPS after a short treatment (10 min) was a rapid release of 2030% of the LPS from the outer membrane and the subsequent sensitization of the bacteria to the anionic detergent sodium dodecyl sulphate.39 The same authors also showed that 20-lysine sensitized smooth E. coli and S. typhimurium strains to hydrophobic antibiotics by a factor of 100.40 In both studies the polymer was not found to be bactericidal.
Our results on the effect of conjugate structure on the PDI efficiency are in accordance with many other reports concerning the use of PDI to kill Gram-negative bacteria. It has been found that the efficacy of a PS in sensitizing Gram-negative bacteria to PDI is related to the charge on the PS itself.11,13 Meso-substituted cationic porphyrins were efficient PSs of Gram-negative bacteria such as Vibrio anguillarum and E. coli after incubation for 5 min.11 In another study, the authors suggested that it was the positive charge that promoted the binding of the porphyrin to the outer membrane, inducing a limited damage that favoured the penetration of the PS.12 They also showed that the photosensitizing activity of cationic porphyrins toward Gram-negative bacteria was inhibited by their incorporation into liposomes.12 Gram-negative bacteria E. coli and P. aeruginosa could be photoinactivated when illuminated in the presence of a cationic water-soluble zinc pyridinium phthalocyanine (PPC) for 30 min but not by illumination in the presence of a neutral tetra-diethanolamine phthalocyanine or a negatively charged tetra-sulphonated phthalocyanine.13 These workers showed recently41 that in-cubation of E. coli cells with a cationic phthalocyanine in the dark caused alterations in the outer membrane permeability barrier and increased the uptake of hydrophobic compounds, with little effect seen with hydrophilic compounds. Addition of Mg2+ to the medium before incubation of the cells with the PS prevented these alterations in the outer membrane permeability barrier and also prevented the PDI of E. coli. PDI of Gram-negative species has been achieved using non-cationic PS in the presence of membrane-disorganizing polycations such as polymyxin B nonapeptide42 or Tris-EDTA, which is also known to disorganize the outer membrane permeability barrier.15
However, there are other reports of PDI of Gram-negative bacteria in which it is clear that the PS does not have to penetrate the bacterium to be effective, or indeed even come into contact with the cells.4346 According to these reports, if singlet oxygen can be generated in sufficient quantities near to the bacterial outer membrane it will be able to diffuse into the cell to inflict damage on vital structures.4346 In one set of studies the bacteria were separated from the PS by a layer of moist air, and singlet oxygen in the gas phase was generated and allowed to diffuse across the gap before contacting the bacteria. Gram-negative species were harder to kill than Gram-positive species, and the intracellular content of carotenoids was found to protect the bacteria from photoinactivation.44 In another study, the PS rose bengal was bound covalently to small polystyrene beads thus preventing the dye from penetrating the bacteria.45 Other workers have bound PS covalently to monoclonal antibodies that recognize and bind to cell surface antigens expressed on P. aeruginosa, and demonstrated specific killing after illumination.46 It is very unlikely that covalent antibody-bound PS could penetrate the outer membrane.
How can these two conflicting sets of findings on the necessity for PS penetration into Gram-negative bacteria be reconciled? One possibility is that singlet oxygen can indeed diffuse into bacteria, producing fatal damage if it is produced at the outer surface or in solution in close proximity. The diffusion distance of singlet oxygen in solution has been estimated to be c. 50 nm,47 and that would indeed allow the molecule to diffuse across the outer membrane to the cytoplasmic membrane. The failure of the 8-lysine conjugate (which had the highest binding to E. coli) to produce any killing must mean that the reactive species produced on illumination were unable to diffuse inwardly to sensitive sites on the cytoplasmic membrane. The fact that this conjugate was the best at killing S. aureus shows that the molecule has not undergone any conformational change that has rendered the PS no longer photoactive. This is presumptive evidence that the reactive species responsible for cell death in the case of Gram-negative species is not singlet oxygen produced by the Type II photoprocess, but rather free radicals and electron transfer processes typical of the Type I photoprocess. In agreement with this hypothesis is a recent report48 in which the PDI of a range of oral pathogenic bacteria was studied using ce6 conjugated to a pL chain with five lysines. It was found that this conjugate efficiently killed a range of Gram-positive species and several Gram-negative anaerobes in the presence of 2.5 mM EDTA and 662 nm light. However, there may be considerable differences between anaerobic and aerobic species in this respect,49 with anaerobes such as P. gingivalis showing sensitivity to both atmospheric oxygen and exogenous reactive oxygen species.50 After completion of the experiments described in the present work, a report by Polo et al.51 appeared reporting similar results. These workers used pLs with two sizes (either 14 kDa or 1530 kDa mean molecular weights) conjugated to porphycene PS and tested their PDI efficacy against E. coli and S. aureus. They found that 1 µM PS equivalent of both conjugates and 27 J/cm2 white light killed S. aureus efficiently, but only the large pL conjugate killed E. coli. The concentration of the small pL conjugate needed to be raised to 10 µM PS equivalent to kill E. coli. They did not examine the uptake of the conjugates by the bacteria and left the conjugates in the bacterial suspension during illumination.
Most reports that discuss the PDI of bacteria propose that the lethal event is damage to the cytoplasmic membrane.52 This damage has been shown to allow vital constituents to leak out into the medium.53 Although damage to DNA has been shown to occur after PDI this is not thought to be the main cause of cell death.54 Deinococcus radiodurans, which has highly efficient DNA repair mechanisms, was still found to be susceptible to PDI.52,55 In agreement with these reports, our data from transmission electron microscopy show that bacterial killing coincides with the appearance of cavities within the cytoplasm, presumably caused by a combination of damage to the membrane and chromosomal changes including DNA condensation.
We have shown previously that both epithelial and endothelial cells accumulate sufficient ce6 from charged pL conjugates to enable efficient photoinactivation to take place.33 However, in mammalian cells this process of uptake is time dependent due to the necessity of these macromolecules being actively internalized via endocytosis. Furthermore, the sparing of HCPC-1 cells in that study under conditions that killed both Gram-positive and Gram-negative bacteria was probably due to the short incubation time.
We have carried out preliminary studies using cationic pL-ce6 conjugates and red light to treat animal models of infected wounds.56 Although it was necessary to use higher concentrations of PS and higher light doses than were required in vitro, it was possible to obtain several logs of bacterial killing in vivo.56 Experiments are underway to define further the utility of this method of PDI of pathogenic bacteria as a means of treating wound and burn infections.
![]() |
Acknowledgements |
---|
![]() |
Footnotes |
---|
![]() |
References |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
2 . Wilson, M. (1993). Photolysis of oral bacteria and its potential use in the treatment of caries and periodontal disease. Journal of Applied Bacteriology 75, 299306.[ISI][Medline]
3 . Carre, V., Gaud, O., Sylvain, I., Bourdon, O., Spiro, M., Blais, J. et al. (1999). Fungicidal properties of meso-arylglycosylporphyrins: influence of sugar substituents on photoinduced damage in the yeast Saccharomyces cerevisiae. Journal of Photochemistry and Photobiology B 48, 5762.[ISI]
4 . Paardekooper, M., Van Gompel, A. E., Van Steveninck, J. & Van den Broek, P. J. (1995). The effect of photodynamic treatment of yeast with the sensitizer chloroaluminum phthalocyanine on various cellular parameters. Photochemistry and Photobiology 62, 5617.[ISI][Medline]
5 . Perlin, M., Mao, J. C., Otis, E. R., Shipkowitz, N. L. & Duff, R. G. (1987). Photodynamic inactivation of influenza and herpes viruses by hematoporphyrin. Antiviral Research 7, 4351.[ISI][Medline]
6 . Ben-Hur, E., Hoeben, R. C., Van Ormondt, H., Dubbelman, T. M. & Van Steveninck, J. (1992). Photodynamic inactivation of retroviruses by phthalocyanines: the effects of sulphonation, metal ligand and fluoride. Journal of Photochemistry and Photobiology B 13, 14552.[ISI]
7 . Bertoloni, G., Salvato, B., DallAcqua, M., Vazzoler, M. & Jori, G. (1984). Hematoporphyrin-sensitized photoinactivation of Streptococcus faecalis. Photochemistry and Photobiology 39, 8116.[ISI][Medline]
8 . Malik, Z., Ladan, H., Nitzan, Y. & Ehrenberg, B. (1990). The bactericidal activity of a deuteroporphyrinhemin mixture on Gram-positive bacteria. A microbiological and spectroscopic study. Journal of Photochemistry and Photobiology B 6, 41930.[ISI]
9 . Bertoloni, G., Rossi, F., Valduga, G., Jori, G., Ali, H. & van Lier, J. E. (1992). Photosensitizing activity of water- and lipid-soluble phthalocyanines on prokaryotic and eukaryotic microbial cells. Microbios 71, 3346.[ISI][Medline]
10 . Malik, Z., Ladan, H. & Nitzan, Y. (1992). Photodynamic inactivation of Gram-negative bacteria: problems and possible solutions. Journal of Photochemistry and Photobiology B 14, 2626.[ISI]
11 . Merchat, M., Bertolini, G., Giacomini, P., Villanueva, A. & Jori, G. (1996). Meso-substituted cationic porphyrins as efficient photosensitizers of Gram-positive and Gram-negative bacteria. Journal of Photochemistry and Photobiology B 32, 1537.[ISI]
12 . Merchat, M., Spikes, J. D., Bertoloni, G. & Jori, G. (1996). Studies on the mechanism of bacteria photosensitization by meso-substituted cationic porphyrins. Journal of Photochemistry and Photobiology B 35, 14957.[ISI]
13 . Minnock, A., Vernon, D. I., Schofield, J., Griffiths, J., Parish, J. H. & Brown, S. B. (1996). Photoinactivation of bacteria. Use of a cationic water-soluble zinc phthalocyanine to photoinactivate both Gram-negative and Gram-positive bacteria. Journal of Photochemistry and Photobiology B 32, 15964.[ISI]
14 . Nitzan, Y., Gutterman, M., Malik, Z. & Ehrenberg, B. (1992). Inactivation of Gram-negative bacteria by photosensitized porphyrins. Photochemistry and Photobiology 55, 8996.[ISI][Medline]
15 . Bertoloni, G., Rossi, F., Valduga, G., Jori, G. & van Lier, J. (1990). Photosensitizing activity of water- and lipid-soluble phthalocyanines on Escherichia coli. FEMS Microbiology Letters 59, 14955.[Medline]
16 . Wainwright, M., Phoenix, D. A., Laycock, S. L., Wareing, D. R. & Wright, P. A. (1998). Photobactericidal activity of phenothiazinium dyes against methicillin-resistant strains of Staphylococcus aureus. FEMS Microbiology Letters 160, 17781.[ISI][Medline]
17 . Sarkar, S. & Wilson, M. (1993). Lethal photosensitization of bacteria in subgingival plaque from patients with chronic periodontitis. Journal of Periodontal Research 28, 20410.[ISI][Medline]
18 . Millson, C. E., Wilson, M., MacRobert, A. J. & Bown, S. G. (1996). Ex-vivo treatment of gastric Helicobacter infection by photodynamic therapy. Journal of Photochemistry and Photobiology B 32, 5965.[ISI]
19 . Groisman, E. A. (1994). How bacteria resist killing by host-defense peptides. Trends in Microbiology 2, 4449.[Medline]
20 . Vaara, M. (1992). Agents that increase the permeability of the outer membrane. Microbiological Reviews 56, 395411.[Abstract]
21 . Vaara, M. (1991). The outer membrane permeability-increasing action of linear analogues of polymyxin B nonapeptide. Drugs Experimental and Clinical Research 17, 43743.[ISI]
22 . Hansen, L. T. & Gill, T. A. (2000). Solubility and antimicrobial efficacy of protamine on Listeria monocytogenes and Escherichia coli as influenced by pH. Journal of Applied Microbiology 88, 104955.[ISI][Medline]
23 . Christensen, B., Fink, J., Merrifield, R. B. & Mauzerall, D. (1988). Channel-forming properties of cecropins and related model compounds incorporated into planar lipid membranes. Proceedings of the National Academy of Sciences, USA 85, 50726.[Abstract]
24 . Duclohier, H., Molle, G. & Spach, G. (1989). Antimicrobial peptide magainin I from Xenopus skin forms anion-permeable channels in planar lipid bilayers. Biophysical Journal 56, 101721.[Abstract]
25 . Ganz, T. (1994). Biosynthesis of defensins and other antimicrobial peptides. Ciba Foundation Symposium 186, 6271.[ISI][Medline]
26 . Gallis, B., Mehl, J., Prickett, K. S., Martin, J. A., Merriam, J., March, C. J. et al. (1989). Antimicrobial activity of synthetic bactenecin. Biotechnology Therapeutics 1, 33546.[Medline]
27 . Weiss, J., Elsbach, P., Shu, C., Castillo, J., Grinna, L., Horwitz, A. et al. (1992). Human bactericidal/permeability-increasing protein and a recombinant NH2-terminal fragment cause killing of serum-resistant Gram-negative bacteria in whole blood and inhibit tumor necrosis factor release induced by the bacteria. Journal of Clinical Investigation 90, 112230.[ISI][Medline]
28 . Tompkins, G. R., ONeill, M. M., Cafarella, T. G. & Germaine, G. R. (1991). Inhibition of bactericidal and bacteriolytic activities of poly-D-lysine and lysozyme by chitotriose and ferric iron. Infection and Immunity 59, 65564.[ISI][Medline]
29 . Helander, I. M., Alakomi, H. L., Latva-Kala, K. & Koski, P. (1997). Polyethyleneimine is an effective permeabilizer of Gram-negative bacteria. Microbiology 143, 31939.[Abstract]
30
.
Soukos, N. S., Ximenez-Fyvie, L. A., Hamblin, M. R., Socransky, S. S. & Hasan, T. (1998). Targeted antimicrobial photochemotherapy. Antimicrobial Agents and Chemotherapy 42, 2595601.
31 . Markwell, M. A., Haas, S. M., Bieber, L. L. & Tolbert, N. E. (1978). A modification of the Lowry procedure to simplify protein determination in membrane and lipoprotein samples. Analytical Biochemistry 87, 20610.[ISI][Medline]
32 . Jett, B. D., Hatter, K. L., Huycke, M. M. & Gilmore, M. S. (1997). Simplified agar plate method for quantifying viable bacteria. Biotechniques 23, 64850.[ISI][Medline]
33 . Soukos, N. S., Hamblin, M. R. & Hasan, T. (1997). The effect of charge on cellular uptake and phototoxicity of polylysine chlorin e6 conjugates. Photochemistry and Photobiology 65, 7239.[ISI][Medline]
34 . Livermore, D. M. (1990). Antibiotic uptake and transport by bacteria. Scandinavian Journal of Infectious Diseases 74, Suppl., 1522.
35 . Sara, M. & Sleytr, U. B. (1987). Molecular sieving through S layers of Bacillus stearothermophilus strains. Journal of Bacteriology 169, 40928.[ISI][Medline]
36 . Hancock, R. E., Farmer, S. W., Li, Z. S. & Poole, K. (1991). Interaction of aminoglycosides with the outer membranes and purified lipopolysaccharide and OmpF porin of Escherichia coli. Antimicrobial Agents and Chemotherapy 35, 130914.[ISI][Medline]
37 . Saberwal, G. & Nagaraj, R. (1994). Cell-lytic and antibacterial peptides that act by perturbing the barrier function of membranes: facets of their conformational features, structurefunction correlations and membrane-perturbing abilities. Biochimica et Biophysica Acta 1197, 10931.[ISI][Medline]
38 . Vaara, M. (1990). The effect of oligolysines Lys-3, Lys-4, and Lys-5 on the outer membrane permeability of Pseudomonas aeruginosa. FEMS Microbiology Letters 55, 159.[ISI][Medline]
39 . Vaara, M. & Vaara, T. (1983). Polycations as outer membrane-disorganizing agents. Antimicrobial Agents and Chemotherapy 24, 11422.[ISI][Medline]
40 . Vaara, M. & Vaara, T. (1983). Polycations sensitize enteric bacteria to antibiotics. Antimicrobial Agents and Chemotherapy 24, 10713.[ISI][Medline]
41
.
Minnock, A., Vernon, D. I., Schofield, J., Griffiths, J., Parish, J. H. & Brown, S. B. (2000). Mechanism of uptake of a cationic water-soluble pyridinium zinc phthalocyanine across the outer membrane of Escherichia coli. Antimicrobial Agents and Chemotherapy 44, 5227.
42 . Nitzan, Y., Dror, R., Ladan, H., Malik, Z., Kimel, S. & Gottfried, V. (1995). Structureactivity relationship of porphines for photoinactivation of bacteria. Photochemistry and Photobiology 62, 3427.[ISI][Medline]
43 . Dahl, T. A., Midden, W. R. & Hartman, P. E. (1987). Pure singlet oxygen cytotoxicity for bacteria. Photochemistry and Photobiology 46, 34552.[ISI][Medline]
44 . Dahl, T. A., Midden, W. R. & Hartman, P. E. (1989). Comparison of killing of Gram-negative and Gram-positive bacteria by pure singlet oxygen. Journal of Bacteriology 171, 218894.[ISI][Medline]
45 . Bezman, S. A., Burtis, P. A., Izod, T. P. & Thayer, M. A. (1978). Photodynamic inactivation of E. coli by rose bengal immobilized on polystyrene beads. Photochemistry and Photobiology 28, 3259.[ISI][Medline]
46 . Friedberg, J. S., Tompkins, R. G., Rakestraw, S. L., Warren, S. W., Fischman, A. J. & Yarmush, M. L. (1991). Antibody-targeted photolysis. Bacteriocidal effects of Sn (IV) chlorin e6-dextran-monoclonal antibody conjugates. Annals of the New York Academy of Sciences 618, 38393.[Abstract]
47 . Ochsner, M. (1997). Photophysical and photobiological processes in the photodynamic therapy of tumours. Journal of Photochemistry and Photobiology B 39, 118.[ISI]
48
.
Rovaldi, C. R., Pievsky, A., Sole, N. A., Friden, P. M., Rothstein, D. M. & Spacciapoli, P. (2000). Photoactive porphyrin derivative with broad-spectrum activity against oral pathogens in vitro. Antimicrobial Agents and Chemotherapy 44, 33647.
49
.
Jenney, F. E., Jr, Verhagen, M. F., Cui, X. & Adams, M. W. (1999). Anaerobic microbes: oxygen detoxification without superoxide dismutase. Science 286, 3069.
50
.
Lynch, M. C. & Kuramitsu, H. K. (1999). Role of superoxide dismutase activity in the physiology of Porphyromonas gingivalis. Infection and Immunity 67, 336775.
51 . Polo, L., Segalla, A., Bertoloni, G., Jori, G., Schaffner, K. & Reddi, E. (2000). Polylysineporphycene conjugates as efficient photosensitizers for the inactivation of microbial pathogens. Journal of Photochemistry and Photobiology B 59, 1528.[ISI]
52 . Schafer, M., Schmitz, C., Facius, R., Horneck, G., Milow, B., Funken, K. H. et al. (2000). Systematic study of parameters influencing the action of Rose Bengal with visible light on bacterial cells: comparison between the biological effect and singlet-oxygen production. Photochemistry and Photobiology 71, 51423.[ISI][Medline]
53 . Malik, Z., Babushkin, T., Sher, S., Hanania, J., Ladan, H., Nitzan, Y. et al. (1993). Collapse of K+ and ionic balance during photodynamic inactivation of leukemic cells, erythrocytes and Staphylococcus aureus. International Journal of Biochemistry 25, 1399406.[ISI][Medline]
54 . Menezes, S., Capella, M. A. & Caldas, L. R. (1990). Photodynamic action of methylene blue: repair and mutation in Escherichia coli. Journal of Photochemistry and Photobiology B 5, 50517.[ISI]
55 . Schafer, M., Schmitz, C. & Horneck, G. (1998). High sensitivity of Deinococcus radiodurans to photodynamically-produced singlet oxygen. International Journal of Radiation Biology 74, 24953.[ISI][Medline]
56 . Hamblin, M. R., ODonnell, D. A., Murthy, N., Contag, C. H. & Hasan, T. (2002). Rapid control of wound infections by targeted photodynamic therapy monitored by in vivo bioluminescence imaging. Photochemistry and Photobiology 75, 517.[ISI][Medline]