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

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

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

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

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

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-

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,

)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-

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

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

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

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

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

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

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

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

`
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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