Báo cáo khoa học: Comparison of human RNase 3 and RNase 7 bactericidal action at the Gram-negative and Gram-positive bacterial cell wall pot

Comparison of human RNase 3 and RNase 7 bactericidal
action 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:
(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.
FEBS Journal 277 (2010) 1713–1725 ª 2010 The Authors Journal compilation ª 2010 FEBS 1713
including 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 evolutionary 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
RNase 3 RNase 7
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
( />and molecular representations were drawn
using
PYMOL (DeLano Scientific, Palo Alto,
CA, USA, ⁄ ).
RNase 3 and RNase 7 bactericidal activity M. Torrent et al.
1714 FEBS Journal 277 (2010) 1713–1725 ª 2010 The Authors Journal compilation ª 2010 FEBS
in 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 (BacLightÔ; 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 ED
50
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. aureus cells 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 filaments 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
⁄
NaH
2
PO
4
) buffer, pH 7.5, and serially diluted from 10 lM to 0.2 lM.
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.
M. Torrent et al. RNase 3 and RNase 7 bactericidal activity
FEBS Journal 277 (2010) 1713–1725 ª 2010 The Authors Journal compilation ª 2010 FEBS 1715
activity was detected in the presence of S. aureus cells,
nor for RNase 7 with the two tested strains, even with
a10lm 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.
Protein E. coli S. aureus
ED
50
(min) Survival (%) Remaining CFUs (%) ED
50
(min) Survival (%) Remaining CFUs (%)
RNase 3 35 ± 1 12.0 ± 0.8 7 ± 4* 60 ± 2 20.1 ± 0.8 22 ± 3*
RNase 7 56 ± 4 17 ± 2 13 ± 3* 56 ± 2 13 ± 1 14 ± 2*
*P < 0.05 (Student’s t-test).
A
B
C
D
E
F
10 min
60 min
120 min
0 min
120 min
RNase 7RNase 3
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.
RNase 3 and RNase 7 bactericidal activity M. Torrent et al.
1716 FEBS Journal 277 (2010) 1713–1725 ª 2010 The Authors Journal compilation ª 2010 FEBS
ribonuclease 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. aureus is 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 DiSC
3
(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.
M. Torrent et al. RNase 3 and RNase 7 bactericidal activity
FEBS Journal 277 (2010) 1713–1725 ª 2010 The Authors Journal compilation ª 2010 FEBS 1717
The 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 activity
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 K
d
value of 2 · 10
)8
m 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
)7
m) [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 lM in 5 mM 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 soluble 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 lLof5mM Hepes-KOH (pH 7.5) and the free and bound fractions
were quantified.
RNase 3 and RNase 7 bactericidal activity M. Torrent et al.
1718 FEBS Journal 277 (2010) 1713–1725 ª 2010 The Authors Journal compilation ª 2010 FEBS
treatment [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
E. coli S. aureus
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.
M. Torrent et al. RNase 3 and RNase 7 bactericidal activity
FEBS Journal 277 (2010) 1713–1725 ª 2010 The Authors Journal compilation ª 2010 FEBS 1719
observed (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).
RNase 3 and RNase 7 bactericidal activity M. Torrent et al.
1720 FEBS Journal 277 (2010) 1713–1725 ª 2010 The Authors Journal compilation ª 2010 FEBS
The 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)a mino)
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
)1
protein 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 (Na
2
HPO
4
⁄ -
NaH
2
PO
4
) buffer (pH 7.5) and serially diluted from 10 lm
to 0.2 lm. Bacteria were incubated at 37 °C overnight in
LB broth and diluted to give approximately 5 · 10
5
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 37 °C 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. coli and S. aureus cells were grown at 37 °C to the mid-
exponential phase (D
600
= 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 (D
600
= 0.2) was mixed
with 5 lm of RNase 3 or 7 and the fluorescence intensity
M. Torrent et al. RNase 3 and RNase 7 bactericidal activity
FEBS Journal 277 (2010) 1713–1725 ª 2010 The Authors Journal compilation ª 2010 FEBS 1721
was 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. ED
50
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
37 °C to the mid-exponential phase (D
600
= 0.6), centri-
fuged at 5000 g for 2 min and resuspended in Tris-HCl
buffer, 0.15 m NaCl (pH 7.5) until D
600
of 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 (D
600
= 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
4 °C 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 4 °C 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)in5mm 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 (ED
50
)
were calculated.
SEM
E. coli and S. aureus cell cultures of 1 mL were grown at
37 °C to the mid-exponential phase (D
600
= 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 scanning electron microscope
RNase 3 and RNase 7 bactericidal activity M. Torrent et al.
1722 FEBS Journal 277 (2010) 1713–1725 ª 2010 The Authors Journal compilation ª 2010 FEBS
(Hitachi, Tokyo, Japan) and a secondary electron image of
cells for topography contrast was collected at several mag-
nifications. A total of ten micrographs were collected at
random for each condition, and the number of isolated cells
and aggregates was registered.
Confocal microscopy
Experiments were carried out in a plate-coverslide system.
Five hundred microlitres of E. coli or S. aureus bacteria
(D
600
= 0.4) were mixed with 40 lLof60lm to achieve a
final concentration of 5 lm of RNase 3 or 7, and images
were immediately recorded. RNase A was used in all cases

as a negative control. Bacteria were pre-stained using
the syto 9 ⁄ propidium iodide 1 : 1 mix provided in the
Live ⁄ Dead staining kit. Syto 9 is a DNA green fluorescent
dye that diffuses thorough intact cell membranes and propi-
dium iodide comprises a DNA red fluorescent dye that can
only access the nucleic acids of membrane damaged cells,
displacing the DNA bound syto 9. The method allows the
labelling of intact viable cells and membrane compromised
cells, which are labelled in green and red respectively,
referred to as live and dead cells [42]. Confocal images of
the bacteria were captured using a laser scanning confocal
microscope (Leica TCS SP2 AOBS equipped with a HCX
PL APO 63, ·1.4 oil immersion objective; Leica Microsys-
tems, Wetzlar, Germany). Syto 9 was excited using a
488 nm argon laser (515–540 nm emission collected) and
propidium iodide was excited using an orange diode (588–
715 nm emission collected). To record the time-lapse experi-
ment, Life Data Mode software (Leica) was used, obtaining
an image every 1 min in a experiment lasting 180 min.
Bacteria cytoplasmic membrane depolarization
assay
Membrane depolarization was assayed by monitoring the
DiSC
3
(5) fluorescence intensity change in response to
changes in transmembrane potential as described previously
[14]. E. coli and S. aureus cells were grown at 37 °C to the
mid-exponential phase and resuspended in 5 mm Hepes-
KOH, 20 m m glucose and 100 mm KCl at pH 7.2 until
D

600
of 0.05 was reached. DiSC
3
(5) was added to a final
concentration of 0.4 lm. Changes in the fluorescence
because of the alteration of the cytoplasmic membrane
potential were continuously monitored at 20 °C at an exci-
tation wavelength of 620 nm and an emission wavelength
of 670 nm. When the dye uptake was maximal, as indicated
by a stable reduction in the fluorescence as a result of
quenching of the accumulated dye in the membrane inte-
rior, protein in 5 mm Hepes-KOH buffer at pH 7.2 was
added at a final tested protein concentration of 4 lm.
Gramicidin D was used as control reference protein. All
conditions were assayed in duplicate. The time required to
reach a stabilized maximum fluorescence reading was
recorded for each condition, and the time required to
achieve half of total membrane depolarization was esti-
mated from the nonlinear regression curve. E. coli cells
were also incubated in the presence of EDTA, allowing loss
of the LPS outer membrane surface layer, as previously
described [14].
Bacteria leakage analysis by activity-staining gels
Activity-staining gels (zymograms) were selected to analyse
bacterial leakage upon incubation with ribonucleases.
E. coli and S. aureus cells were grown at 37 °C to the mid-
exponential phase (D
600
= 0.4) in LB medium, centrifuged
at 5000 g for 5 min, and resuspended in a 10 mm Na

2
HPO
3
buffer at pH 7.2. Cells were incubated with 5 lm of
RNase 3 or 7 and 10 lL aliquots were collected at 1, 2, 3
and 4 h. The aliquots collected were mixed with nonreduc-
ing loading buffer (60 mm Tris-HCl, 10% glycerol, 0.015%
bromophenol blue, 3% SDS, pH 6.8) and analysed for
RNase activity by zymogram on SDS-PAGE (15%) con-
taining 0.6 mgÆmL
)1
of polycytidylic acid as substrate.
After elimination of SDS by incubation with a pH 7.5 solu-
tion consisting of 10 mm Tris-HCl and 20% isopropanol,
the gels were incubated at 25 °C for 90 min in 100 mm
Tris-HCl (pH 7.5). The relative intensity of the areas show-
ing substrate degradation was analysed by densitometry.
Bacterial cell leakage was assessed by monitoring, as a
function of time, the increase of the clearance area corre-
sponding to polynucleotide cleavage by the released bacte-
rial RNase.
Acknowledgements
Confocal microscopy and scanning electron micros-
copy were performed at the Servei de Microsco
`
pia of
the Universitat Auto
`
noma de Barcelona (UAB). We
thank Mo

`
nica Rolda
´
n and Helena Monto
´
n for their
technical support with confocal microscopy, and Fran-
cisca Cardoso, Francesc Bohils and Alejandro Sa
´
nchez
for their assistance with the electron microscopy
samples. Spectrofluorescence and densitometry assays
were performed at the Laboratori d’Ana
`
lisi i Fotodoc-
umentacio
´
, UAB. The work was supported by the
Ministerio de Educacio
´
n y Cultura (grant numbers
BFU2006-15543-C02-01 and BFU2009-09371) and by
the Fundacio
´
La Marato
´
de TV3 (grant number TV3-
031110). M.T. was the recipient of a predoctoral
fellowship from the Generalitat de Catalunya.
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Supporting information
The following supplementary material is available:
Fig. S1. Study of S. aureus viability and population
morphology upon incubation with RNase 3 visualized
by confocal microscopy.
Fig. S2. Study of S. aureus viability and population
morphology upon incubation with RNase 7 visualized
by confocal microscopy.
Fig. S3. Analysis by SDS-PAGE of RNase 3 cell bind-
ing to E. coli and S. aureus cells.
Table S1. RNase 3 and RNase 7 membrane depolar-
ization activities.
Video S1. Register of E. coli viability and population
morphology upon incubation with RNase 3, stained
with the Live ⁄ Dead kit and visualized by confocal
microscopy.
This supplementary material can be found in the
online version of this article.
Please note: As a service to our authors and readers,
this journal provides supporting information supplied
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from supporting information (other than missing files)
should be addressed to the authors.
M. Torrent et al. RNase 3 and RNase 7 bactericidal activity
FEBS Journal 277 (2010) 1713–1725 ª 2010 The Authors Journal compilation ª 2010 FEBS 1725