RNase 3 displays a specific Escherichia coli cell agglutination activity, which is not shared by RNase 7.. We hypothesize that the RNase 3 agglutination activity may depend on its high af
Trang 1action at the Gram-negative and Gram-positive bacterial cell wall
Marc Torrent, Marina Badia, Mohammed Moussaoui, Daniel Sanchez, M Victo`ria Nogue´s and Ester Boix
Departament de Bioquı´mica i Biologia Molecular, Facultat Biocie`ncies, Universitat Auto`noma de Barcelona, Cerdanyola del Valle`s, Spain
Introduction
Human antimicrobial RNase 3 and RNase 7 are
mem-bers of the RNase A superfamily that participate in
the host immune response against pathogen infection
RNase 3 was first identified as an eosinophil secretion
product and named as eosinophil cationic protein
(ECP) ECP is secreted by activated eosinophils during
inflammation and its levels in biological fluids are
con-sidered to be a marker for the diagnosis and
monitor-ing of allergy and eosinophilia disorders [1] Recently,
it was reported that eosinophils can mediate their anti-bacterial effect through the release of cationic granule proteins [2] RNase 7 was first reported as a skin antimicrobial protein [3] and is considered to be one of the main components of the innate immunity first-line protection against infections at the epithelial level [4,5] RNase 7 is expressed in several epithelial tissues,
Keywords
antimicrobial proteins; cell wall; ECP;
immunity; RNase 7
Correspondence
E Boix, Departament de Bioquı´mica i
Biologia Molecular, Facultat de Biocie`ncies,
Universitat Auto`noma de Barcelona, 08193
Cerdanyola del Valle`s, Spain
Fax: +34 93 5811264
Tel: +34 93 5814147
E-mail: ester.boix@uab.cat
(Received 19 November 2009, revised
25 January 2010, accepted 27 January
2010)
doi:10.1111/j.1742-4658.2010.07595.x
The eosinophil cationic protein⁄ RNase 3 and the skin-derived RNase 7 are two human antimicrobial RNases involved in host innate immunity Both belong to the RNase A superfamily and share a high cationicity and a common structural architecture However, they present significant diver-gence at their primary structures, displaying either a high number of Arg
or Lys residues, respectively Previous comparative studies with a mem-brane model revealed two distinct mechanisms of action for lipid bilayer disruption We have now compared their bactericidal activity, identifying some features that confer specificity at the bacterial cell wall level RNase 3 displays a specific Escherichia coli cell agglutination activity, which is not shared by RNase 7 The RNase 3 agglutination process precedes the bacte-rial death and lysis event In turn, RNase 7 can trigger the release of bacterial cell content without inducing any cell aggregation process We hypothesize that the RNase 3 agglutination activity may depend on its high affinity for lipopolysaccharides and the presence of an N-terminal hydro-phobic patch, and thus could facilitate host clearance activity at the infec-tion focus by phagocytic cells The present study suggests that the membrane disruption abilities do not solely explain the protein bacterial target preferences and highlights the key role of antimicrobial action at the bacterial cell wall level An understanding of the interaction between anti-microbial proteins and their target at the bacterial envelope should aid in the design of alternative peptide-derived antibiotics
Abbreviations
CFU, colony-forming unit; ECP, eosinophil cationic protein; MAC, minimal agglutination concentration; PGN, peptidoglycan; SEM, scanning electron microscopy.
Trang 2including skin, gut and the respiratory and
genitouri-nary tracts, and its expression can be induced by
inflammatory agents and bacterial infection [6] Both
RNases display a wide range anti-pathogen activity,
with toxicity being reported against viruses, bacteria,
fungi, protozoans and, in the case of RNase 3, even
helminthic parasites [7] Although both proteins belong
to the RNase A superfamily and have conserved their
catalytic RNase activity [3,8], studies indicate that their
antimicrobial mechanism of action is strongly
depen-dent on their membrane destabilizing mechanism of
action [9,10] The RNase A superfamily includes other
members with antimicrobial properties [7] and recent
evolution studies suggest that the family may have
started with an ancestral antipathogen physiological
function [11,12] Previous experimental data on both
RNases, using lipid vesicles as a membrane model,
revealed that the lipid bilayer disruption event takes
place with a distinct timing [10,13] However, the data
obtained also indicate that mechanic action at the
cytoplasmic membrane does not solely explain the
pro-tein bactericidal properties Therefore, we also
charac-terized RNase 3 activity at the surface of bacteria,
identifying significant differences with respect to its
action on both Gram-negative and Gram-positive
strains A key distinctive feature of RNase 3 is its high
affinity for lipopolysaccharides (LPS) and Escherichia
coli cell agglutination activity [14] Despite the fact
that both RNases show a high cationicity, they share approximately 40% amino acid identity; careful inspec-tion reveals a distinct evoluinspec-tionary pressure that leads
to the accumulation of an unusual number of either Arg (18 Arg out of 133 amino acids) or Lys (18 Lys out of 128 amino acids) at the mature protein (Fig 1) Mutagenesis studies indicated the involvement of positive and aromatic surface-exposed residues for RNase 3 [15] and some surface lysine clusters for RNase 7 [9] On the other hand, a binding domain for heparin in RNase 3 [16] may account for its high affinity for heterosaccharides at the bacterial cell wall Indeed, recent studies using RNase 3-derived peptides revealed a key domain at the protein N-terminus, which retained most of the protein bactericidal activity and a considerable LPS binding capacity [17] More-over, screening of the RNase 3 N-terminal sequence predicts a hydrophobic aggregation patch [9] and an antimicrobial prone sequence [18]
We have now compared the activity of both RNases
at the bacterial cell wall level Although RNase 7 dis-plays remarkable affinity for peptidoglycan (PGN) and LPS at the Gram-positive and Gram-negative outer surface, the very high LPS binding and cell agglutina-tion activities represent a distinctive feature of RNase 3 By contrast, RNase 7 displays a high leakage activity and a high capacity for binding PGN The comparison of both antimicrobial RNases conducted
A
C
B
Fig 1 (A) Ribbon representation of the 3D structures of RNase 3 (1DYT.pdb) [43] and RNase 7 (2HKY.pdb) [9] Molecules are coloured from the N- to C-terminus The active site is marked by a circle (B) Molecular surface representation of RNase 3 and RNase 7 Hydrophobic residues are labelled in grey, cationic residues in blue, anionic residues in red, cysteine residues in yellow, proline residues
in orange and noncharged polar residues in cyan (C) Sequence alignment of RNase 3 and RNase 7 primary sequences Secondary structure elements of RNase 3 are depicted
at the top The sequence alignment was performed using ESPRIPT software (http://espript.ibcp.fr/ESPript/ESPript/) and molecular representations were drawn using PYMOL (DeLano Scientific, Palo Alto,
CA, USA, http://www.pymol.org ⁄ ).
Trang 3in the present study therefore contributes towards
elu-cidating the main determinants of their distinct
poten-tial in vivo anti-pathogen properties
Results
Studies on the bacterial cell viability
We have compared the RNase 3 and RNase 7
antimi-crobial activities with respect to E coli and Staphylococcus
aureus cells, which are representative Gram-negative
and Gram-positive strains Both proteins display
com-parable activity, as indicated by the reduction of
col-ony-forming units (CFUs) as a function of protein
concentration (Fig 2) On the other hand, kinetic
pro-files of bacterial viability show a similar overall pattern,
although there were significant differences in the
respec-tive activities for the two tested strains The bactericidal
activity profiles were monitored by staining of bacteria
with a Live⁄ Dead kit (BacLight; Molecular Probes,
Carlsbad, CA, USA), using syto 9 and propidium
iodide to determine bacterial viability Although syto 9
dye can cross the cytoplasmic membrane and label all
bacterial cells, propidium iodide can only access the
content of membrane damaged cells, competing and
displacing the bound syto 9 Therefore, the integration
of syto 9 and propidium iodide fluorescence provides
an estimate of the percentage viability for monitoring
the kinetics of the bactericidal process (Fig 3)
Although RNase 7 shows a similar live⁄ dead
progres-sion for both studied bacterial species, RNase 3 is
sig-nificantly more active on the E coli population, as
reflected by the ED50 values (Fig 3 and Table 1)
The relative percentage survival, as evaluated by the
viability assay, also correlated with the reduction in the
percentage of remaining CFUs (Table 1)
To determine the morphological changes in bacterial
cell population upon incubation with both RNase 3
and 7, the process was also visualized using confocal
microscopy, where live⁄ dead cells are also labelled with
the syto 9 and propidium iodide dyes, respectively
A careful inspection on the culture population
behaviour by confocal microscopy reveals how
RNase 3 aggregates E coli cells, and how bacterial cell
death is a later event in relation to the aggregation
process (Fig 4) By comparison, RNase 3-treated
S aureuscells display a distinct behaviour, where
bac-terial death takes place at only a slightly lower rate
but without a significant aggregation pattern (Fig S1)
Therefore, we conclude that the results obtained for
RNase 3 indicate that the key bactericidal events take
place at different times First, we observe an
enlarge-ment on the filaenlarge-ments formed by E coli cells The
structures formed (after 10–20 min of incubation) are only stained by syto 9, indicating that these filaments are formed by live bacteria From 30 min onward, the bacterial population stained by propidium iodide is rapidly increased Subsequently, the aggregates begin
to bind propidium iodide and recruit new dead clusters
of bacteria (Video S1) For S aureus, this aggregation mechanism cannot be observed and only an increase in the propidium iodide-stained bacteria is detected Although some small clusters of bacteria can be observed, they are not comparable to the aggregates obtained in the case of E coli For RNase 7, aggrega-tion is neither observed in E coli, nor in S aureus (Figs 4 and S2)
To quantify the bacterial aggregation ability, the minimal agglutination concentration (MAC) was calcu-lated, with an estimated value of 1.5 lm for RNase 3 activity with E coli cells, whereas no agglutination
Fig 2 Remaining CFUs after exposure of bacterial cultures to (A)
E coli and (B) S aureus The response is registered as a function
of the protein concentration RNase 3 (triangles) and RNase 7 (squares) were dissolved in 10 m M sodium phosphate (Na 2 HPO 4 ⁄ NaH2PO4) buffer, pH 7.5, and serially diluted from 10 l M to 0.2 l M
In each assay, protein solutions were added to each dilution of bacteria, incubated for 4 h, plated in Petri dishes and the colonies counted after overnight incubation.
Trang 4activity was detected in the presence of S aureus cells,
nor for RNase 7 with the two tested strains, even with
a 10 lm protein concentration The results obtained
show that RNase 7 lacks the ability to agglutinate bacteria but retains bactericidal activity
To better understand the correlation between aggregation and bacterial leakage, the release of cell content was monitored using activity staining gels (Fig 5) With this technique, the endogenous bacterial
Fig 3 Study of bacterial viability kinetics for (A) RNase 3 and (B)
RNase 7 Cell viability for Gram-positive S aureus (filled squares)
and Gram-negative E coli (filled circles) was analysed using syto 9
(for live bacteria) and propidium iodide (for dead bacteria) An
aliquot of 1 mL of exponential phase cells was incubated with 5 l M
of each protein Duplicates were performed for each condition.
Table 1 Kinetic analysis on the antimicrobial activity of RNases 3 and 7 using the Live ⁄ Dead bacterial viability kit as described in the Materi-als and methods One millilitre of exponential phase cells was incubated with 5 l M of protein during a total period of 150 min ED 50 (mea-sured as the time needed to achieve a 50% decrease in live bacteria) and percentage survival were calculated by exponential fitting to the data presented in Fig 3 The percentage of remaining CFUs is also indicated for each condition Values are the average of three replicates.
ED50(min) Survival (%) Remaining CFUs (%) ED50(min) Survival (%) Remaining CFUs (%)
*P < 0.05 (Student’s t-test).
A
B
C
D
E
F
10 min
60 min
120 min
0 min
120 min
Fig 4 Study of E coli viability and population morphology visual-ized by confocal microscopy E coli cells (A) before protein addi-tion; (B–D) after 5 l M of RNase 3 at 10 min (B), 1 h (C) and 2 h (D); and (E, F) after adding 5 l M of RNase 7 at 0 and 2 h, respectively Bacterial cells were stained using a 1 : 1 syto 9 ⁄ propidium iodide mixture The left-hand panels correspond to the propidium iodide-stained cells (dead cells), excited using an orange diode The cen-tral panels correspond to the syto 9-stained cells (live cells), excited using a 488 nm argon laser The right-hand panels correspond to the overlay of both signals Scale bar = 50 lm.
Trang 5ribonuclease released upon membrane leakage can be
detected and the leakage kinetics can be monitored
The bacterial cells were incubated with 5 lm of each
RNase and aliquots were taken at 1-h intervals For
RNase 3, an important difference between E coli and
S aureusis found Whereas leakage in E coli cells can
be observed as soon as after 1 h of incubation, no
release is detected for S aureus, not even after 4 h of
incubation These results demonstrate that, even
though RNase 3 is able to kill 80% of S aureus cells
after 4 h of incubation, the damage at the membrane
level is insufficient to allow the release of a detectable
amount of endogenous ribonucleases
In the case of RNase 7, both E coli and S aureus
endogenous RNases are released (Fig 5) Nevertheless,
RNase 7 leakage in S aureus cells appears to be
trig-gered later than in E coli cells The activity
corre-sponding to the endogenous ribonucleases that are
released by the bacteria is only registered after 2 h of
incubation
Finally, membrane depolarizing activity was also
studied using the DiSC3(5) marker (Table S1) The
results obtained show that RNase 3 is able to
depo-larize E coli cells more rapidly than S aureus cells
When comparing membrane depolarization activities,
we can observe that ECP easily accesses the
Gram-negative cytoplasmic membrane, without any EDTA
treatment being necessary to destabilize the cell outer
membrane RNase 3 activity on E coli cells is
inde-pendent of EDTA chelation This is not applicable to RNase 7, which has a lower membrane depolarization activity without EDTA treatment On the other hand, RNase 7 appears to alter more easily the S aureus cytoplasmic membrane than RNase 3 The distinct abilities of both RNases to access and alter the cyto-plasmic membrane may reflect their action at the outer envelope level
Studies at the bacterial cell wall The bactericidal activity of both RNases is precluded
by the protein binding to the cells Proteins incubated with both E coli and S aureus cultures are recovered
in the cell pellet fraction (Fig S3) To gain insight on the bactericidal properties of both RNases, binding studies on different elements of the bacterial cell wall were carried out Binding to PGN and LPS has already been studied in detail for RNase 3 [14] The results obtained are now compared with RNase 7 binding affinities The new data (Figs 6 and 7) indicate that RNase 7 can also interact with both Gram-negative and Gram-positive heteropolysaccharides Affinity binding studies on LPS and PGN were com-plemented with scanning electron microscopy (SEM) microscopy to visualize the structural damage induced
by the protein–cell wall interactions (Fig 8)
Binding to LPS was assessed using the Bodipy TR cadaverine marker (Invitrogen, Carlsbad, CA, USA)
A
B
Fig 5 Record of bacterial lysis process by the detection of the release of endogeneous bacterial RNase by activity staining gel (A) The clearance area corresponding to the bacterial RNase substrate degradation is indicated The intensity of the areas showing substrate degradation was analysed by densitometry as described in the Materials and methods The intensity values are referred to the 0 h incubation density area The bacterial lysis activity of RNase 3 (filled symbols) and RNase 7 (empty symbols) on both E coli (triangles) and S aureus (squares) is shown (B) Polycytidylic acid SDS-PAGE (15%) activity staining gel from the time course of E coli cell incubation with RNase 3 Left lanes: control cells; right lanes: cells incubated with 5 l M of RNase 3 at 0, 1, 2, 3 and 4 h.
Trang 6The results obtained show that RNase 3 is able to bind
with higher affinity to LPS compared to RNase 7 In
any case, RNase 7 still retains a high LPS binding
affin-ity because it displays an effective displacement activaffin-ity
similar to that for polymyxin B, a powerful LPS binder, which was selected as a positive control (Fig 6)
We also assessed and compared RNase 7 binding to PGN, the main component of Gram-positive bacteria, with our previous results obtained for RNase 3 [14] Microfluidic gel electrophoresis showed that, after RNase 7 incubation in the presence of S aureus PGN, most of the protein sample is recovered together with the insoluble PGN fraction, as also observed for lyso-zyme, the positive control, and previously for RNase 3 [14] A slight anomalous displacement in the virtual gel
is observed for RNase 7, with a higher apparent molecular weight, as a result of its cationic nature This behaviour is frequently observed for RNase A family members By contrast, BSA, the negative con-trol, does not bind to the PGN fraction and is fully recovered in the supernatant fraction (Fig 7A) Moreover, a PGN binding assay using Alexa fluoro-phor-labelled RNase 7 also indicates a high binding affinity A Kd value of 2· 10)8m was determined using the Scatchard plot as shown in Fig 7B, which is
a value considerably higher than that calculated for RNase 3 (2· 10)7m) [14]
SEM data were previously shown to be useful for assessing bacterial surface damage upon RNase 3
Fig 6 Displacement of LPS-bound Bodipy TR cadaverine by
RNase 7 (triangles), RNase 3 (circles) and polymyxin B (squares);
[LPS]: 10 lgÆmL)1; [BODIPY TR Cadaverine]: 10 l M in 5 m M
He-pes-KOH (pH 7.5).
100.0 75.0 50.0 37.0 25.0 20.0
B A
Fig 7 (A) Analysis by a microfluidic electrophoresis system of the binding of RNase 7 to PGN Lysozyme and BSA were taken as positive and negative controls, respectively, for PGN binding Molecular mass markers are indicated on the left For each protein, the first lane corre-sponds to pellet (P) and the second lane to the supernatant fractions (S) PGN were incubated with each protein and the soluble and insolu-ble fractions were collected as described in the Materials and methods Supernatant represents the soluinsolu-ble fraction, which contains the unbounded protein, whereas the pellet represents the insoluble fraction containing the PGN bound protein (B) Scatchard plot and the corre-sponding binding curve of RNase 7 interaction with PGN RNase 7 labelled with the fluorophor Alexa Fluor 488 at a concentration in the range 0.01–100 n M was incubated in the presence of 0.02 lg PGN in 200 lL of 5 m M Hepes-KOH (pH 7.5) and the free and bound fractions were quantified.
Trang 7treatment [14], where severe damage on E coli cells
and the ability of protein to trigger cell population
agglutination was reported Accordingly, SEM was
used to visualize changes in bacterial cell cultures upon
incubation with RNase 7 The addition of RNase 7 at
a final concentration of 4 lm is unable to induce either
E coli or S aureus cell culture aggregation and all
cells retain their characteristic morphology
Neverthe-less, several blebs can be observed on the bacterial cell
surface in both E coli and S aureus, suggesting that
local cell surface disturbance is taking place (Fig 8)
Discussion
RNases 3 and 7 are the main representatives of the
cytotoxic antimicrobial members of the RNase A
superfamily Both are cationic proteins with a high pI,
and display a broad antimicrobial action against
Gram-positive and Gram-negative strains [6,19–21]
The two RNases present, respectively, a high number
of either Arg or Lys surface-exposed residues (Fig 1)
that may contribute to their distinct bactericidal
mech-anisms of action Previous work revealed that the
RNase bactericidal mechanism was not dependent on
its RNase enzymatic activity but on direct membrane
disruptive action [9,10,15,22] The contribution of
bac-terial wall determinants was also suggested [15] and
recent studies on RNase 3 indicated a high affinity for
bacterial heterosaccharides [14] Indeed, the present
comparative characterization of both the action of
RNase 3 and RNase 7 at the bacterial wall level
revealed some particular features that could explain
their distinct abilities with respect to Gram-negative
and Gram-positive strains We previously compared
the mechanism of action of both RNases on model membranes [10,13] RNase 7 has no significant mem-brane aggregation capacity compared to RNase 3, although it displays a much higher leakage capacity
On the other hand, initial studies on RNase 3 by site-directed mutagenesis indicated that the membrane dis-ruption ability could not solely explain the protein bactericidal properties [15] Indeed, strain selectivity was reported for RNase 7 [3,9]
We have now analysed the time course profile of bacterial cell viability for both RNases (Fig 3) The rapid decay during the first 30 min may reflect a rapid direct lytic process We can differentiate between an initial active exponential growth phase, where the pro-tein may have easy access to the cell membrane during duplication, and a later stage, where protein action at the wall envelope may acquire a critical role On the other hand, the viability assay, performed at a salt concentration close to physiological levels, rejects a mere unspecific electrostatic interaction and provides further corroboration for both proteins retaining their properties in vivo and being regarded as effective anti-microbial agents As noted by Hancock and Sahl [23], many cationic peptides with few hydrophobic residues
at crucial positions are prone to having some antimi-crobial activity at low ionic strength, although the term ‘antimicrobial’ should only be reserved for those that are able to kill microbes under physiological conditions
The results obtained in the present study reveal dis-tinct behaviours not only on lipid bilayers, but also at the bacterial cell wall In both strains, E coli and
S aureus, RNase 7 displays a restricted disturbance causing local blebs, whereas no agglutination is
Fig 8 Scanning electron micrographs of
E coli and S aureus incubated in the
absence (top) and presence (bottom) of
4 l M RNase 7 for 4 h The magnification
scale is indicated at the bottom of each
micrograph.
Trang 8observed (Fig 8) These observations are much
differ-ent from those observed in the case of RNase 3, where
global cell damage has been observed in E coli cells
after complete bacterial agglutination [14]
Interestingly, the in vivo record of the RNase 3
trea-ted E coli culture assessed by confocal microscopy
illustrates how the cells first aggregate but still retain
an intact cytoplasmic membrane Cell death, as
observed by the propidium iodide uptake, is then a
later event (Fig 4 and Video S1) We have further
analysed RNase 3 bacterial agglutination activity and
estimated a MAC of 1.5 lm on E coli cell cultures
Cell agglutination comprises a characteristic feature
that also is reported for other antimicrobial peptides
[24] and proteins, as lectin RNases, which are
amphi-bian members of the RNase A superfamily with a
particular ability for binding heterosaccharides [25]
In turn, RNase 7 could follow another bacterial
pro-cess The ability to induce the bacterial cell content, as
assayed by activity-staining gel analysis, has shown
that, in S aureus, RNase 7 presents an important
leakage activity, whereas no significant activity is
detected for RNase 3 at the assayed conditions
(Fig 5) This fact may be explained by the higher
capacity of RNase 7 to cause leakage of membranes at
low concentrations These effects are in good
agree-ment with the results observed in model membranes,
where RNase 7 is able to trigger leakage at a lower
protein : lipid ratio before any aggregation event takes
place, suggesting a local membrane disturbance process
[10] Moreover, the higher binding affinity for PGN
displayed by RNase 7 may also partially account for
the higher membrane depolarization activity observed
against the S aureus strain (Table S1) RNase 7 was
previously reported to display a particularly high
bac-tericidal activity for the Gram-positive
Enterococ-cus faecium [3] Our membrane depolarizing assays
confirm a distinct mechanism of action for both
RNases on each of the two tested strains Mainly for
Gram-negative cells, RNase 3 does not require EDTA
pretreatment EDTA pretreatment would sequester the
divalent cations that hold LPS together and secure the
outer membrane structure The higher affinity of
RNase 3 for LPS (Fig 6) could by itself facilitate
outer membrane disturbance and access to the
cyto-plasmic membrane RNase 7 displays a similar
capac-ity for depolarizing cell membranes, as observed in
RNase 3, when E coli cells are pretreated with EDTA,
thus suggesting that the main differences may be
restricted to the bacterial outer barrier
These results confirm that the capacity to bind
bac-terial cell wall structures is of special importance for
the antimicrobial properties of both RNases, as also
reported for other antimicrobial proteins and peptides [26,27] If we compare the sequences and 3D structures available for both RNases (Fig 1), we can identify some of the features that may account for the specific ability of RNase 3 to aggregate both lipid vesicles and bacterial cells Scanning of both RNases with aggreg-scan software [28] reveals a distinct aggregation pro-file, in particular at the N-terminal zone In the case of RNase 3, we can observe a hydrophobic patch in one side of the molecule, surrounded by polar residues Indeed, a hydrophobic patch at the RNase 3 N-termi-nus that retains most of the protein antimicrobial activity, and may be responsible for the protein vesicle aggregation ability, was recently characterized by syn-thetic-derived peptides in our laboratory [17] Bacteria agglutinating efficiency was also correlated with the presence of hydrophobic patches for de novo designed antimicrobial peptides In the case of RNase 7, no hydrophobic patches on the protein surface can be observed The protein cationicity, as a result of the high number of lysines present in the structure, is dis-tributed uniformly on the protein surface The absence
of hydrophobic patches may be responsible for the lack of agglutinating capacity of RNase 7
Although both RNases contain a high number of cationic residues, the bias on either Arg or Lys con-tent (18 Arg for RNase 3 and 18 Lys for RNase 7) suggests that the cationicity of both proteins has been acquired independently during their evolution A comparison with other RNase A family members indicates that most Lys residues are retained in the RNase A lineage group that includes RNases 6, 7 and 8 [29] Phylogenetic studies suggest the recent divergence of RNase 7 and RNase 8 as a result of a duplication event [29] However, no homologues were identified in rodents [12] as described for the RNase2⁄ RNase 3 group, where members with antimi-crobial activity were reported in both rat and mouse
In turn, RNase 3 acquired many Arg residues during its divergence from RNase 2 [12,29] However, a com-parison of antimicrobial RNases suggests that local positive clusters, rather than their overall pI, are key for protein bactericidal activities [30,31] For example,
a comparison of the primary sequences for fish, chicken and human antimicrobial RNases revealed a distinct Lys⁄ Arg ratio but a similar total number of positive residues [30]
On the other hand, arginine residues are implied in carbohydrate binding proteins because they display hydrogen bonding between the guanidinium group and sulphates or phosphates [32,33] This fact may explain the higher binding affinity of RNase 3 for LPS (Fig 6)
Trang 9The tissue distribution of both RNases also suggests
some functional differences Whereas RNase 3 is
mostly present in eosinophils and, to a less extent, in
other cells of the immune system (e.g neutrophils and
basophils) [34,35], RNase 7 is expressed in multiple
somatic tissues, especially the skin, where it is
described as a major antimicrobial agent [3,6]
Although both RNases are secreted, they may respond
to distinct challenges RNase 3 is stored in secretion
granules and is depleted at the site of inflammation
where these cells are recruited [36] RNase 7 represents
one of the major contributors to the antimicrobial
activity involved in first-line host defence at the human
skin barrier [37] In the skin, basal RNase 7 secretion
is detected but mRNA overexpression is observed as a
result of bacterial challenge [37] A correlation between
a dysfunction in antimicrobial protein expression at
the skin level during dermatitis and a predisposition to
skin infections also highlights their contribution to a
host defence role [38–40]
In conclusion, in the present study, we have shown
that RNase 3 and RNase 7 have particular
antimicro-bial activities that are modulated by their action at the
bacterial cell wall We observed that RNase 7 displays
a mechanism based on local membrane disturbance, in
contrast to RNase 3 that demonstrated global action
Accordingly, we have shown that RNase 3 displays an
E coli agglutinating activity (not shared by RNase 7),
which would probably be dependent on both the
pres-ence of a hydrophobic patch and the capacity of the
protein to bind LPS
An understanding of the molecular mechanism that
is responsible for the high binding affinity of
antimi-crobial protein for unique heterosaccharide structures
at the bacterial envelope would also contribute to the
development of new peptide-derived antibiotics, which
would overcome the increasing emergence of antibiotic
resistant strains
Materials and methods
Materials
Bodipy TR cadaverine, BC
[5-(((4-(4,4-difluoro-5-(2-thienyl)-4-bora-3a,4a-diaza-s-indacene-3-yl)phe-noxy)acetyl)amino)
pentylamine, hydrochloride], 3,3-dipropylthiacarbocyanine
[DiSC3(5)], Gramicidin D, Alexa Fluor 488 protein
label-ling kit and the Live⁄ Dead bacterial viability kit were all
purchased from Molecular Probes (Eugene, OR, USA)
LPSs from E coli serotype 0111:B4, Polymyxin B sulfate,
PGN from S aureus, polycytidylic acid and lysozyme from
chicken egg white were purchased from Sigma-Aldrich
(St Louis, MO, USA) E coli BL21DE3 (Novagen, Madison,
WI, USA) and S aureus 502 A (ATCC, Rockville, MD, USA) strains were used PD-10 columns were purchased from GE Healthcare (Milwaukee, WI, USA)
Expression and purification of recombinant RNase 3 and RNase 7
Wild-type RNase 3 was expressed using a synthetic gene for human coding sequence RNase 7 was expressed start-ing from a cDNA subcloned in the pET11c plasmid vector Protein expression in E coli BL21(DE3) strain, folding of the protein from inclusion bodies, and the purification steps, were carried out as described previously [8,10]
Fluorescent labelling of proteins
RNases were labelled with the Alexa Fluor 488 fluorophor,
in accordance with the manufacturer’s instructions To 0.5 mL of a 2 mgÆmL)1protein solution in NaCl⁄ Pi, 50 lL
of 1 m sodium bicarbonate (pH 8.3) was added The pro-tein was incubated for 1 h at room temperature with the reactive dye, with stirring, in accordance with the manufac-turer’s instructions The labelled protein was separated from the free dye by a PD-10 desalting column
Antibacterial activity
Antimicrobial activity was calculated by assessing the num-ber of CFUs as a function of protein concentration Values were averaged from two independent experiments per-formed in triplicate for each protein concentration Proteins were dissolved in 10 mm sodium phosphate (Na2HPO4⁄ -NaH2PO4) buffer (pH 7.5) and serially diluted from 10 lm
to 0.2 lm Bacteria were incubated at 37C overnight in
LB broth and diluted to give approximately 5· 105
CFUÆmL)1 In each assay, protein solutions were added to each dilution of bacteria, incubated for 4 h, and samples were plated on Petri dishes and incubated at 37C over-night The number of CFUs in each Petri dish was counted and the average values were represented in a semi-logarith-mic plot
Bacterial viability
Kinetics of bacterial survival were carried out using the Live⁄ Dead bacterial viability kit in accordance with the manufacturer’s instructions Bacteria were stained using a syto 9⁄ propidium iodide 1 : 1 mix as provided with the kit
E coliand S aureus cells were grown at 37C to the mid-exponential phase (D600= 0.4), centrifuged at 5000 g for
5 min and resuspended in a 0.75% NaCl solution in accor-dance with the manufacturer’s instructions One millilitre of stained E coli or S aureus bacteria (D600= 0.2) was mixed with 5 lm of RNase 3 or 7 and the fluorescence intensity
Trang 10was continuously measured using a Cary Eclipse
Spectroflu-orimeter (Varian Inc., Palo Alto, CA, USA) RNase A was
used in all cases as a negative control The excitation
wave-length was 470 nm and the emission was recorded in the
range 490–700 nm To calculate bacterial viability, the
sig-nal in the range 510–540 nm was integrated to obtain the
syto 9 signal (live bacteria) and from 620–650 nm to obtain
the propidium iodide signal (dead bacteria) Then, the
per-centage of live bacteria was represented as a function of
time ED50 was calculated by fitting the data to a simple
exponential decay function
Agglutination activity
Agglutination activity was evaluated by calculating the
MAC An aliquot of 5 mL of E coli cells was grown at
37C to the mid-exponential phase (D600= 0.6),
centri-fuged at 5000 g for 2 min and resuspended in Tris-HCl
buffer, 0.15 m NaCl (pH 7.5) until D600of 10 was reached
An aliquot of 200 lL of the bacterial suspension was
incu-bated in microtitre plates with an increasing protein
con-centration at 0.1 and 0.5 lm intervals up to 10 lm and left
overnight at room temperature The aggregation behaviour
was observed by visual inspection and checked with a
bin-ocular microscope at ·50 magnification The agglutinating
activity is expressed as the minimum agglutinating
concen-tration of the sample tested, corresponding to the first
con-dition where bacterial aggregates are visible by the naked
eye, as described previously [41]
Protein binding to bacterial cells
RNase 3 was incubated at 5 lm with E coli bacterial cells
grown to the exponential phase (D600= 0.6) in 1 mL of
NaCl⁄ Pi buffer at 37 C for 1 h After centrifugation at
13 000 g, proteins from the pellet were extracted with
electrophoresis loading buffer Supernatant fractions
were lyophilized and dissolved in loading buffer Samples
were analysed by SDS-PAGE (15%) and Coomassie blue
staining
Affinity binding assay for PGN
Protein binding to PGN was first analysed by
electrophore-sis as described previously [14] PGN at 0.4 mgÆmL)1 in
10 mm Tris-HCl (pH 7.5) was incubated with the protein at
a protein⁄ PGN ratio of 1 : 20 (w ⁄ w) Samples were kept at
4C for 2 h with gentle mixing and centrifuged at 13 000 g
for 15 min to separate the soluble and insoluble fractions
Lysozyme and BSA were chosen as positive and negative
controls, respectively Samples were resuspended directly in
the electrophoresis loading buffer and evaluated using an
Experion automated microfluidic electrophoresis system
(Bio-Rad, Hercules, CA, USA)
Protein affinity to PGN was calculated using a fluores-cence-based method, employing a microtitre plate as described previously [14] Protein labelled with the fluoro-phor Alexa Fluor 488 was incubated with insoluble PGN Proteins at different concentrations, in the range 1–100 nm, were incubated in the presence of 0.02 lg of peptidoglycans
in a 5 mm Hepes buffer at pH 7.5 in a final volume of
200 lL The reaction mixture was kept at 4C for 2 h with gentle shaking Next, the remaining soluble protein was removed from the insoluble PGN fraction by a centrifuga-tion step at 13 000 g for 30 min and quantified with Victor 3 (Perkin-Elmer, Boston, MA, USA)
Affinity binding assay for LPS
LPS binding was assessed using the fluorescent probe Bodipy TR cadaverine as described previously [14] Briefly, the displacement assay was performed by the addition of 1–2 lL aliquots of a solution of Polymyxin B, RNase 3, RNase 7 or RNase A to 1 mL of a continuously stirred mixture of LPS (10 lgÆmL)1) and Bodipy TR cadaverine (10 lm) in 5 mm Hepes buffer at pH 7.5 Fluorescence measurements were performed on a Cary Eclipse spec-trofluorimeter The BC excitation wavelength was 580 nm and the emission wavelength was 620 nm The excitation slit was set at 2.5 nm and the emission slit was set at
20 nm Final values correspond to an average of four repli-cates and were the mean of a 0.3 s continuous measure-ment Quantitative effective displacement values (ED50) were calculated
SEM
E coli and S aureus cell cultures of 1 mL were grown at
37C to the mid-exponential phase (D600= 0.4) and incu-bated with 4 lm RNase 3 or RNase 7 in NaCl⁄ Pi at room temperature Aliquots were taken up to 4 h of incubation and were prepared for analysis by SEM, as described previ-ously [14] The cell suspensions were fixed with 2.5% gluter-aldehyde in 100 mm Na-cacodylate buffer (pH 7.4) for 2 h
at room temperature Next, the cells were pelleted, a drop
of each suspension was transferred to a nucleopore filter, which was kept in a hydrated chamber for 30 min allowing the cells to adhere, and then washed to remove the gluteral-dehyde, and resuspended in the same 100 mm Na-cacody-late buffer at pH 7.4 Attached cells were post-fixed by immersing the filters in 1% osmium tetroxide in cacodylate buffer for 30 min, rinsed in the same buffer, and dehy-drated in ethanol in ascending percentage concentrations [31, 70, 90 (·2) and 100 (·2)] for 15 min each The filters were mounted on aluminum stubs and coated with gold-palladium in a sputter coater (K550; Emitech, East Grinsted, UK) The filters were viewed at 15 kV accelerat-ing voltage in a Hitachi S-570 scannaccelerat-ing electron microscope