In situ probing of newly synthesized peptidoglycan in live bacteria with fluorescent D-amino acids

Article abstract of DOI:10.1002/anie.201206749

Tracking a bug's life: Peptidoglycan (PG) of diverse bacteria is labeled by exploiting the tolerance of cells for incorporating different non-natural D-amino acids. These nontoxic D-amino acids preferably label the sites of active PG synthesis, thereby enabling fine spatiotemporal tracking of cell-wall dynamics in phylogenetically and morphologically diverse bacteria. HCC=7-hydroxycoumarin, NBD=7-nitrobenzofurazan, TAMRA= carboxytetramethylrhodamine. Copyright

Full text of DOI:10.1002/anie.201206749

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Chemie
DOI: 10.1002/anie.201206749
Fluorescence Imaging
In Situ Probing of Newly Synthesized Peptidoglycan in Live Bacteria
with Fluorescent d-Amino Acids**
Erkin Kuru, H. Velocity Hughes, Pamela J. Brown, Edward Hall, Srinivas Tekkam, Felipe Cava,
Miguel A. de Pedro, Yves V. Brun,* and Michael S. VanNieuwenhze*
Bacterial growth is controlled by the peptidoglycan (PG) cell
wall, a rigid and essential structure composed of glycan
strands cross-linked by d-amino acid (DAA)-containing short
peptides, the biosynthesis machinery of which is a high-value
target for antibiotics.[1] Despite the incredible importance of
this polymer, our knowledge of PG dynamics has been
severely hampered by the lack of a global strategy for direct
imaging of sites of PG synthesis in live cells. Limitations of
current labeling methods, such as toxic effects and poor
membrane permeability of the probes, have limited their
applicability to only a small set of bacterial species.[2]
Commonly, these methods are labor-intensive and their
sensitivity suffers from their indirect and multiple-step
nature. To overcome these limitations, we sought a chemical
biology approach that would enable the rapid and covalent
incorporation and detection of a fluorescently derivatized PG
component during cell wall synthesis in real time, in a wide
range of bacterial species, and in live cells.
a modular and simple synthetic scheme, and generated the
fluorescent amino acids HADA (emission maximum 450 nm,
blue) and NADA (emission maximum 538 nm, green;
Scheme 1 and Figure S2 in the Supporting Information).
Growth of the phylogenetically diverse model species
Escherichia coli, Agrobacterium tumefaciens, and Bacillus
subtilis in the presence of these fluorescent d-amino acids
(FDAAs) for as little as one generation (Figure S1a in the
Given the recently established role of the PG biosynthetic
machinery in incorporating various natural DAAs in the PG
of diverse bacteria,[3] we hypothesized that the mechanisms[3b]
for DAA incorporation in PG are common to the bacterial
domain but still highly selective, and that molecules with
various functionalities could be incorporated into the sites of
new PG synthesis if these molecules are attached to a DAA
backbone. To test this hypothesis, we attached the relatively
small fluorophores, 7-hydroxycoumarin-3-carboxylic acid
(HCC-OH) and 4-chloro-7-nitrobenzofurazan (NBD-Cl), to
a d-amino acid backbone, 3-amino-d-alanine, by using
Scheme 1. Structures of the fluorescent d-amino acids HADA, NADA,
FDL, and TDL with the fluorescent group shown in the color of the
fluorophore (blue, green, and red) and the “clickable” d-amino acids
EDA and ADA.
[*] E. Kuru, H. V. Hughes, Dr. P. J. Brown, E. Hall, S. Tekkam,
Prof. Y. V. Brun, Prof. M. S. VanNieuwenhze
Indiana University
Supporting Information) resulted in strong peripheral and
septal labeling of entire cell populations (Figure 1 and
Figure S3a in the Supporting Information, upper panels)
without affecting growth rate (Figure S3b in the Supporting
Information). Neither of the fluorescent l-amino acids
(FLAAs, prepared from 3-amino-l-alanine, Figure S2c in
the Supporting Information) resulted in significant labeling
(Figure S3c), thus indicating that labeling is specific to the d-
enantiomers. The labeling was exclusive to viable cells treated
with the FDAAs and was not the result of nonspecific
interaction of FDAAs with the PG (Figure S4 in the
Supporting Information). Moreover, incorporation did not
occur into teichoic acids for B. subtilis as indicated by
identical labeling of wild-type and a DdltA mutant that does
not incorporate d-alanine into its teichoic acids (Figure S5a,b
in the Supporting Information).
Bloomington, IN 47405 (USA)
E-mail: ybrun@indiana.edu
mvannieu@indiana.edu
Prof. F. Cava, Prof. M. A. de Pedro
Universidad Autonoma de Madrid, Campus de Cantoblanco
Madrid, 28049 (Spain)
[**] We thank David Kysela for discussions and critical reading of the
manuscript, Bill Turner and Alvin Kalinda for their help in
simplifying the synthesis of FDAAs and Prof. Patricia Foster and the
members of her laboratory for their support during the initial stages
of the project. M.S.V. was funded by National Institutes of Health
(NIH) grant AI 059327, Y.V.B. by NIH grant GM051986, M.A.P. by
Ministry of Education and Science (Spain) grant BFU2006-04574
and by the Fundaciꢀn Ramꢀn Areces, F.C. by MICINN (Spain) grant
RYC-2010-06241, and P.J.B. by NIH National Research Service
Award AI072992.
Retained fluorescence on the purified sacculi (Figure 1
and Figure S3a in the Supporting Information, lower panels)
Supporting information for this article is available on the WWW
under http://dx.doi.org/10.1002/anie.201206749.
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unlabeled muropeptides at FDAA-specific absorption wave-
lengths (Figure 2b). MS/MS analyses of FDAA-modified
muropeptides in B. subtilis indicated that FDAAs were
exclusively incorporated in the fifth position of the stem
peptide (Figure 2d, Figure S7a,d and Table S1 in the Sup-
porting Information). Interestingly, in a DdacA mutant of
B. subtilis, the fraction of labeled muropeptides and the
fluorescent signal were substantially higher than in wild-type
B. subtilis. The diminshed signal intensity in wild-type cells is
likely due to the d,d-carboxypeptidase activity of DacA
(Figure S5c,d in the Supporting Information). In contrast, the
detectable incorporation was solely at the fourth position in
E. coli and A. tumefaciens (Figure 2d and Figure S7 in the
Supporting Information). These results are in agreement with
the known sites of incorporation of various natural DAAs in
these species and suggest that, similar to DAAs, FDAAs
incorporate mainly through periplasmic exchange reactions
with the muropeptides; these reactions are catalyzed either by
d,d-transpeptidases, for example, in B. subtilis, or by l,d-
transpeptidases, for example, in E. coli and A. tumefa-
ciens.[3b,4] This DAA-like behavior together with the ease of
fluorescent detection make FDAAs a strong alternative to
radioactive probes for studying activities of PG synthesis
Figure 1. Long labeling pulses with HADA uniformly label PG in live
cells of E. coli, B. subtilis, and A. tumefaciens. The FDAA fluorescence is
retained in isolated sacculi, which are also stained with the N-acetyl-
glucosamine (GlcNAc) specific WGA lectin conjugated to Alexa Fluor
594 (red). Scale bars: 2 mm.
demonstrated that the labeling of PG by the FDAAs was
covalent. HPLC analyses of muropeptides isolated from
labeled cells (Figure 2a–c, Figures S6 and S7 in the Support-
ing Information) revealed that 0.2–2.8% of total muropep-
tides were modified (Figure 2c), which is sufficient for
detection in various experiments while avoiding possible
toxicity issues that could result from abundant incorporation.
Significantly, the FDAA-specific peaks, which were absent in
samples treated with FLAAs, could be distinguished from
enzymes in vitro[3c] and in vivo[5]
.
In contrast to current approach-
es,[2b,f] pulse–chase experiments with
HADA allowed the real-time tracking
of new PG incorporation during
growth by time–lapse microscopy. In
E. coli and B. subtilis DdacA (Fig-
ure 3a and Movies 1 and 2 in the
Supporting Information), the polar
caps retained the HADA signal, but
the signal from the lateral walls dis-
persed as the cells grew; this observa-
tion is in agreement with reports of cell
wall growth along the length of the
lateral walls.[2f,5]
Strikingly, short labeling times (2–
8% of doubling time) using E. coli and
B. subtilis DdacA with HADA resulted
in preferential localization of the
signal at the septal plane of predivi-
sional cells and in punctate patterns on
the lateral walls of elongating cells
(Figure 4). Super-resolution microsco-
py of E. coli revealed reticulated
hoop-like patterns of HADA labeling
around the lateral wall (Figure 3b and
Movie 3 in the Supporting Informa-
tion); these patterns support the find-
ing of locations with a high density of
PG in the side walls.[6] This ability of
FDAAs to resolve insertion of new PG
enables the first direct detection of
zones of PG synthesis in a structured
pattern rather than in a random pat-
tern in E. coli; this result is consistent
with recent reports describing the
Figure 2. FDAA incorporation into the stem peptide of the PG subunit. a) Schematic representa-
tion showing the muramylpentapeptide precursor unit. b) HPLC detection of modified muropep-
tides in E. coli incubated with HADA or HALA and NADA or NALA. Samples were monitored
using a dual-wavelength UV monitor set for general muropeptide detection and for FDAA-specific
wavelengths. Peaks HEC-1 and NEC-1 correspond to the HADA- or NADA-modified muropeptides
in E. coli and were further characterized by electrospray ionization MS/MS (ESI–MS/MS). The
numbers 1–3 in the chromatograms assign the peaks to the peptides of the respective lengths.
c) Percentage of FDAA incorporation into the total muropeptides varies among different bacteria
as revealed by HPLC analysis. d) MS/MS analyses show that FDAAs exclusively incorporate into
the 4th position of muropeptides in E. coli and A. tumefaciens and the 5th position in B. subtilis.
See also Figures S6,7 and Table S1 in the Supporting Information. MurNAc=N-acetylmuramic
acid; m-DAP=meso-diaminopimelic acid.
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Figure 3. FDAAs label diverse bacterial growth patterns. a) Time-lapse
microscopy of HADA-labeled E. coli and B. subtilis DdacA cells imaged
during growth on LB agarose pads (LB=Luria Bertani). b,c) Super-
resolution microscopy of E. coli (b) and A. tumefaciens (c) after short
pulses with HADA. d) Super-resolution microscopy of S. aureus cells
after a short pulse with HADA. Autofluorescence is shown in red.
e) Triple labeling of A. tumefaciens with HADA (blue), EDA (clicked
with red sulfo-Cy3-azide), and NADA (green). f) Triple labeling of S.
venezuelae with NADA (green), TdL (red), and HADA (blue). Arrows in
the triple-labeling panels indicate the sequence of labeling. White scale
bars: 2 mm; red scale bars: 1 mm.
Figure 4. Short pulses of HADA label distinct modes of growth in
diverse bacteria. Strains were labeled for ca. 2–8% of the doubling
time: E. coli (30 s), A. tumefaciens (2 min), B. subtilis DdacA (30 s),
S. aureus (2 min), L. lactis (2 min), S. pneumoniae (4 min), C. crescentus
(5 min), Synechocystis sp. PCC 6803 (1 h), S. venezuelae (2 min), B. con-
glomeratum (8 min), B. phytofirmans (20 min), V. Spinosum (10 min).
Scale bars: 2 mm.
representing diverse phyla and modes of growth were briefly
incubated with FDAAs, we observed strong labeling at the
sites of cell division in actively dividing cells (Figure 4 and
Figure S1b in the Supporting Information). This septal probe
incorporation was the sole mode in Synechocystis sp. PCC
6803, Lactococcus lactis, and Staphylococcus aureus as
previously described.[8] Super-resolution microscopy of Staph-
ylococcus aureus further highlighted the different stages of
these constricting septal rings (Figure 3d and Movie 5 in the
Supporting Information). Labeling of Streptococcus pneumo-
niae occurred in single or split equatorial rings depending on
the length of the cell, with peripheral labeling between the
split rings as described.[8] Labeling of Streptomyces venezuelae
was predominantly apical as reported,[9] with some weak
labeling of vegetative septa and lateral walls, thus suggesting
circumferential movement of the cell wall elongation machi-
nery.[7] Short labeling times with A. tumefaciens, the growth of
which occurs predominantly from a single pole and the site of
cell division while the mother cell remains inert,[4] resulted in
polar and septal labeling. Super-resolution fluorescence
microscopy of labeled cells further enhanced the spatial
resolution of the site of active PG synthesis (Figure 3c and
Movie 4 in the Supporting Information).
PG labeling in three evolutionarily distant species sug-
gested that FDAAs could specifically label the active site of
PG synthesis across the entire bacterial domain. When species
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a low but continuous lateral PG synthesis. In Caulobacter
crescentus, labeling occurred at the sites of septal elongation,
lateral elongation, and stalk synthesis, as reported else-
where.[10]
carrying the complementary functional group.[14] Similar to
FDAAs, these bioorthogonal DAAs, but not the l-enantio-
mer control amino acid ELA, labeled both E. coli and
B. subtilis cells when captured by commercially available
azido/alkyne fluorophores (Figures S2c,d and S9d,e in the
Supporting Information).
Furthermore, custom d-amino acids containing different
colored fluorophores can be used sequentially to enable
“virtual time-lapse microscopy.” Since addition of each new
probe indicates the location and extent of PG synthetic
activity during the respective labeling periods, this approach
provides a chronological account of shifts in PG synthesis of
individual cells over time. This powerful method is exempli-
fied here through click chemistry in gram-negative A. tume-
faciens (Figure S1a in the Supporting Information and Fig-
ure 3e), or by the use of TDL in gram-positive S. venezuelae
(Figure 3 f).
In contrast to the bacterial protein-making machineryꢀs
inherent bias against non-natural amino acids,[15] here we have
shown that taxonomically diverse bacteria display a remark-
able specific tolerance for incorporating d-amino acids with
different sizes and functionalities into PG. By exploiting this
tolerance we developed a rapid, nontoxic, and universal
method for real-time tracking of PG at sites of active synthesis
for the first time. Moreover, the combined utilization of two
or more probes permits a temporal resolution that was never
before achieved in cell wall growth studies. We expect that in
combination with fluorescent fusion proteins, mutational
analysis, and chemical perturbations, this methodology will
allow a comprehensive analysis of the regulation and
coordination of bacterial growth. Furthermore, the tolerance
for DAAs with substantial sizes or with bioorthogonal
handles will enable selective and specific modification of
bacterial cell surfaces with various functionalities, thus paving
the way for development of DAA-based bacteria-specific
diagnostic or therapeutic probes. Finally, when cells labeled
with FDAAs are hybridized with fluorescent in situ hybrid-
ization (FISH) probes, this approach will allow the culture-
independent and concurrent measurement of growth (with
FDAAs) and taxonomy (with FISH) and the response of
bacteria to varying conditions in medical or environmental
microbiomes.
Bacteria exhibit a myriad of growth patterns that provide
selective advantages in the environment.[11] The strong
correlation between FDAA labeling and previously inferred
regions of new PG synthesis in diverse model species has
a number of important implications. First, FDAA labeling
marks the sites of active PG synthesis and therefore provides
a long-sought broadly applicable tool to study the spatial
dynamics of PG synthesis. Second, these results establish for
the first time that the enzymes responsible for DAA
incorporation in the PG are associated with active growth
sites and that different areas on the surface of the sacculus are
not equally accessible to these enzymes. Finally, FDAAs can
be used to discover the growth modes of previously unchar-
acterized taxa. For example, our results show that Burkhol-
deria phytofirmans exhibits polar and midcell PG synthesis,
that Brachybacterium conglomeratum exhibits prominent
peripheral PG synthesis in addition to seemingly alternating
perpendicular division planes, and that Verrucomicrobium
spinosum exhibits strong peripheral PG synthesis and asym-
metric septal labeling (Figure 4).
The efficient label incorporation in all the bacteria studied
thus far also suggests that FDAA incorporation, and there-
fore DAA incorporation, is common to the bacterial domain
and FDAAs can thus be used to analyze natural bacterial
populations, thereby providing a convenient and quick
standard to measure bacterial activity and to probe the
diversity of growth modes in complex microbiomes.[11]
Indeed, labeling times with FDAAs as short as two hours
revealed diverse modes of growth in saliva and freshwater
samples in situ, but did not label dead cells as suggested by the
strong correlation with live–dead staining (Figures S4b and S8
in the Supporting Information).
Encouraged by the efficiency of FDAAs, we sought to
increase our toolkit for PG detection and modification using
differently functionalized non-natural d-amino acids. By
following a similar approach, we derivatized the brighter
and more versatile core fluorophore, fluorescein (emission
maximum ca. 515 nm, green), and its analogue, carboxy-
tetramethylrhodamine (TAMRA, emission maximum ca.
565 nm, red),[12] with d-lysine to separate the bulky fluoro-
phore from the DAA backbone, thereby generating FDAAs
FdL and TDL (Scheme 1 and Figure S2a in the Supporting
Information). Incubation of both E. coli and B. subtilis cells
with FDL (536 Da) resulted in patterns similar to those with
NADA, although labeling of B. subtilis was stronger than
E. coli, presumably owing to reduced permeability of the
E. coli outer membrane to molecules larger than approx-
imately 500 Da[13] (Figure S9a–c in the Supporting Informa-
tion). Indeed, the larger TDL (560 Da) did not label E. coli
cells, but labeling of B. subtilis was prominent and gave
patterns similar to other FDAAs (Figure S9a,b in the
Supporting Information). To expand the toolkit further, we
used “clickable” d-amino acids, namely ethynyl-d-alanine
(EDA) or azido-d-alanine (ADA; Scheme 1), which can be
specifically captured by using click chemistry by any molecule
Received: August 20, 2012
Published online: October 10, 2012
Keywords: bacteria · biosensors · d-amino acids ·
.
fluorescent probes · peptidoglycan
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