its research

1. Review
Chemical Biology Tools
for Examining the Bacterial Cell Wall
Ashley R. Brown,1 Rebecca A. Gordon,2,3 Stephen N. Hyland,1 M. Sloan Siegrist,2,3 and Catherine L. Grimes1,4,*
1Department of Chemistry and Biochemistry, University of Delaware, Newark, DE 19716, USA
2Department of Microbiology, University of Massachusetts, Amherst, MA 01003-9298, USA
3Molecular and Cellular Biology Graduate Program, University of Massachusetts, Amherst 01003-9298, USA
4Department of Biological Sciences, University of Delaware, Newark, DE 19716, USA
*Correspondence: cgrimes@udel.edu
https://doi.org/10.1016/j.chembiol.2020.07.024
SUMMARY
Bacteria surround themselves with cell walls to maintain cell rigidity and protect against environmental in-
sults. Here we review chemical and biochemical techniques employed to study bacterial cell wall biogenesis.
Recent advances including the ability to isolate critical intermediates, metabolic approaches for probe incor-
poration, and isotopic labeling techniques have provided critical insight into the biochemistry of cell walls.
Fundamental manuscripts that have used these techniques to discover cell wall-interacting proteins, flip-
pases, and cell wall stoichiometry are discussed in detail. The review highlights that these powerful methods
and techniques have exciting potential to identify and characterize new targets for antibiotic development.
INTRODUCTION
The bacterial cell wall is arguably just as important to human
health as it is to the survival of the bacterium. The complex poly-
mers that comprise the cell wall provide bacteria with strength
and a barrier to the outside world, allowing them to thrive in a
multitude of environments, including the human body. Humans
have taken advantage of natural product antibiotics, often pro-
duced by bacteria themselves, to target bacterial polymers,
yielding some of the most widely used antibiotics to date (Cho-
pra and Roberts, 2001). In this review, we focus on understand-
ing the bacterial cell wall in the context of the threat that antibiotic
resistance poses to society. With the announcement that many
major pharmaceutical companies will no longer fund research
programs in this critical area (Hu, 2018), the onus falls on curious,
determined academics to identify new targets for antibiotics and
new opportunities to combat resistance. Here we highlight
recently reported chemical and biochemical approaches to
study bacterial cell wall biosynthesis and maintenance both in
the laboratory and in the clinic, focusing on the use of antibiotics.
In some cases, antibiotics are part of the toolkit that enables bio-
logical insight. In other cases, the biological insights gleaned
from applying the tools enable (or have the potential to enable)
target discovery. Studies of the bacterial cell wall and the antibi-
otics that corrupt it are iterative and promote both mechanistic
insights and translational applications.
THE TARGET AND ITS BASICS
Scientists have used small molecules to study bacteria since the
19th
century when Christian Gram treated cells with crystal violet
and realized that bacteria could be canonically divided into
two general classes: Gram-positive and Gram-negative
(Gram, 1884). In the present day, sophisticated tools exist to
visualize the cell wall and to dissect its composition and biosyn-
thesis at the molecular level (Hsu et al., 2019; Kocaoglu and Carl-
son, 2016; Radkov et al., 2018; Siegrist et al., 2015; Taguchi
et al., 2019a). Here we will review a subset of the new methods,
but first offer a brief introduction to cell walls, noting the many
recent, detailed reviews on these structures (e.g., Radkov
et al., 2018) and their biosynthesis and maintenance (Taguchi
et al., 2019a).
The bacterial cell envelope is a complex structure that pro-
vides protection from the external environment, maintains cell
shape, and provides resistance to chemical, physical, and bio-
logical damage (Figure 1A; Vollmer et al., 2008). Nearly all bacte-
rial envelopes have a peptidoglycan (PG) cell wall layer lying just
outside the plasma membrane. PG biosynthesis is a highly
conserved process in bacteria, starting with UDP-N-acetyl-
glucosamine (UDP-GlcNAc) conversion into UDP-N-acetyl-mur-
amic acid (UDP-MurNAc), as the first committed step in PG syn-
thesis (Figure 1B). The subsequent steps involve the addition of
amino acids (commonly L-Ala, D-g-Glu, L-Lys [Gram-positive] or
meso-diaminopimelic acid [mDAP; Gram-negative and myco-
bacteria], and D-Ala-D-Ala) to the UDP-MurNAc lactate moiety.
Variation exists within the stem peptide depending on species.
For example, Mycobacterium leprae can utilize Gly in place of
L-Ala at the one position (Draper et al., 1987), many Gram-posi-
tive bacteria and mycobacteria amidate the second position to
post-biosynthetically generate D-g-Gln, and spirochetes include
L-Orn at the third position (Schleifer and Kandler, 1972; Vollmer
et al., 2008). These and additional variations are highlighted in
Vollmer et al. (2008). Stem peptide ligation is followed by the
transfer of this PG building block to phosphate polyprenyl to
make Lipid I. GlcNAc is added to Lipid I to make Lipid II as the
final PG precursor. Lipid II is subsequently flipped across the
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2. membrane by the MurJ flippase (Ruiz, 2008; Sham et al., 2014)
as the complete PG subunit. Class A penicillin-binding proteins
(aPBPs), L,D-transpeptidases, and shape, elongation, division,
and sporulation (SEDS) proteins in complex with class B PBPs
(bPBPs) assemble the PG through two different enzymatic reac-
tions, transglycosylation and transpeptidation (Cho et al., 2016;
Emami et al., 2017; Meeske et al., 2016; Taguchi et al., 2019b).
PG transglycosylases (TGs) link the sugar backbone of the PG
subunit to the next unit through a b-1,4 glycosidic linkage
(Vollmer et al., 2008). PG transpeptidases (TPs) most commonly
connect the fourth amino acid of one peptide chain to the third
amino acid (such as mDAP or L-Lys) of an adjacent strand
yielding 3–4 cross-linked PG (Vollmer and Seligman, 2010).
However, there are more variations that feature diverse connec-
tivity—3-3, 2-4, and 1-3 linkages—and bridge lengths across
species. While the biosynthesis of Lipid II is generally conserved
across bacteria, the polymerization of the monomer marks the
beginning of the diversification process (Egan et al., 2020; Typas
et al., 2011). Depending on the activity and the protein-protein in-
teractors of the TGs/TPs, a variety of shapes are formed (Do
et al., 2020; Salama, 2020). For example, in H. Pylori, the TG
and TP activity is spatially and temporally regulated along the
cell wall, which greatly influences the shape. This diversity of
Gram-positive and Gram-negative cells is further enhanced by
the inclusion of lipids in the cell wall.
In Gram-positive bacteria, the PG layer is significantly thicker
than that of Gram-negative (Vollmer and Seligman, 2010) and
features lipoteichoic and wall teichoic acids (WTAs) anchored
to the cytoplasmic membrane and the MurNAc 6-OH, respec-
tively. The WTA is assembled largely on the cytoplasmic face.
Upon delivery to the extracellular surface, it is anchored to PG
by the family of LytR-CpsA-Psr (LCP) enzymes (Kawai et al.,
2011; Li et al., 2020; Schaefer et al., 2017). Teichoic acid (TA) el-
ements are essential to the virulence of pathogenic bacteria.
They permit bacterial cell adhesion to host cells, in addition to
controlling cell wall remodeling by autolysins (Brown et al., 2013).
Gram-negative bacteria have a thinner PG but have additional
structural support and protection due to the asymmetric outer
membrane consisting of phospholipids and lipopolysaccharides
(LPS) (Rojas et al., 2018). LPS is made of three components: lipid
A, an oligosaccharide core, and the O-antigen. Toward the end
of the lipid A construction, the core oligosaccharide is added,
making the lipid oligosaccharide intermediate. The O-antigen is
synthesized separately and bound to lipooligosaccharide
(LOS), to make LPS, before transport to the outer membrane
(Simpson and Trent, 2019). LPS is not anchored to the PG but
rather inserted into the outer membrane by complex cellular ma-
chinery and hydrophobically adhered via Lipid A.
There is a third class of bacteria that has elements of both
Gram-positives and Gram-negatives; it is sometimes referred
Figure 1. Bacterial Cell Wall Basics
(A) Structural features of the cell wall of Gram-positives, Gram-negatives, and mycobacteria. PG, peptidoglycan; PM, plasma membrane; LPS, lipopolysac-
charide; MM, mycomembrane; OM, outer membrane; AG, arabinogalactan.
(B) Biosynthetic steps of peptidoglycan (PG): PG biosynthesis occurs in 14 unique biochemical steps, starting with UDP-GlcNAc conversion into UDP-MurNAc.
UDP-MurNAc is transferred to the lipid carrier and, subsequently, glycosylated by MurG. This process culminates in Lipid II being flipped across the membrane by
MurJ. Transglycosylases (TGs) and transpeptidases (TPs) incorporate the PG subunits into the growing PG polymer. Ultimately, peptidoglycan is altered through
the addition of small and large molecules post-synthetically that are not represented here, such as wall teichoic acids (WTAs).
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3. to as Gram-indeterminate. M. tuberculosis and other members
of the Corynebacterineae suborder are phylogenetically related
to Gram-positive bacteria, but possess a waxy, relatively imper-
meable outer membrane-like structure (Figure 1A; Alderwick
et al., 2015). The core cell wall has a PG layer of intermediate
thickness that is covalently bound to the branched arabinogalac-
tan (AG). The AG layer acts as a scaffold for a covalently bound,
inner leaflet of mycolic acids. The outer leaflet possesses more
mycolic acids, along with intercalated glycolipids such as treha-
lose monomycolates (TMMs) and trehalose dimycolate (TDM)
(Dulberger et al., 2020). The three classes of bacteria discussed
here contain some similarities in the composition of their cell wall
(i.e., Lipid II; Figure 1B) and differences (i.e., lipid modifications).
It has been proposed that some of the lipid modifications that
help to differentiate Gram-positive, Gram-negative, and Gram-
indeterminate bacteria could provide mechanisms to uniquely
target specific bacteria with antibiotic therapy (Jackson et al.,
2013; Kuhn, 2019).
Bacterial cell walls have historically made an excellent target for
antibiotics. There are multiple ways to kill a bacterial cell, inhibiting
DNA replication and halting protein synthesis (Walsh, 2003). How-
ever, the bacterial cell wall is an especially attractive target
because it is unique to bacteria, meaning that human cells do
not contain the biochemical machinery that is required to build
and maintain it. In addition, the biosynthesis is largely conserved
(Figure 1B), allowing the development of broad-spectrum antibi-
otics. Unfortunately, bacteria have developed resistance to nearly
every cell wall-targeting antibiotic, including the well-known b-lac-
tams, such as penicillin and methicillin, and the antibiotics of ‘‘last
resort,’’ the glycopeptides (Kahne et al., 2005). Modes of resis-
tance can be conferred by, but not limited to, the expression of
insensitive PBPs, production of b-lactamases, activity of efflux
pumps, and alterations to the cell wall (Walsh, 2000). For a more
extensive review of these and other modes of resistance, the au-
thors refer the reader to Walsh (2000). In recent years, there has
been an emergence of multi-drug-resistant bacteria, particularly
in the ESKAPE pathogens (Enterococcus faecium, Staphylo-
coccus aureus, Klebsiella pneumoniae, Acinetobacter baumanii,
Pseudomonas aeruginosa, and Enterobacter species) (Boucher
et al., 2009; Santajit and Indrawattana, 2016). These bacteria,
together with tuberculosis (Bloom and Murray, 1992; Porter and
McAdam, 1994) and gonorrhea (Unemo and Shafer, 2014), pose
a viable threat with a lack of new antimicrobial compounds
(CDC, 2017). Within 1 year after the first clinical use of a natural
or synthetic antibiotic, resistance can develop (Walsh, 2003).
Therefore, there is a continuous demand for the development of
novel antibiotics via rational design, high-throughput screening ef-
forts, and medicinal chemistry campaigns.
In order to identify new bactericidal, cell wall-acting com-
pounds, peptidoglycan biosynthetic pathways (Figure 1) that
are less understood and not currently targeted by mainstream
antibiotics must be interrogated with appropriate tools. Fluores-
cent probes have permitted the close study of bacterial struc-
tures, and more recently probes such as fluorescent D-amino
acids (FDAAs) have enabled methods to screen for novel antibi-
otics and to identify their mode of action (Culp et al., 2020). There
are many excellent reviews that highlight direct imaging of bac-
terial cell walls to study the structure, biosynthesis, and dy-
namics of this polymer (Kocaoglu and Carlson, 2016; Radkov
et al., 2018; Siegrist et al., 2015; Taguchi et al., 2019a). Here
we focus on recent work that merges chemical and biochemical
methods including immunoblotting, photo-crosslinking, spec-
troscopy, and radiolabeling to probe PG biosynthetic pathways
and composition that can help inform the development of novel
antimicrobial therapeutics.
USING ANTIBIOTICS TO ISOLATE CELL WALL
INTERMEDIATES
In order to study bacterial cell wall biosynthetic enzymes, one
must have access to PG precursor substrates. A common strat-
egy is to use bacterial cell wall inhibitors such as vancomycin and
moenomycin to cause a buildup of intermediates. Such a tactic
was used by Strominger and Park in the 1950s to identify the
UDP-MurNAc intermediate, known as Park’s nucleotide, using
penicillin (Figure 1B) (Park, 1952; Strominger et al., 1959). The
lipid-linked precursor, Lipid II, was long sought in PG biosyn-
thesis (Figure 1B) as it is difficult to synthesize, and its scarcity
made in vitro studies of PG polymerization nearly impossible
(Lazar and Walker, 2002; Ye et al., 2001). Lipid II is the result of
the b-1,4 glycosylation of Lipid I by MurG with the monosaccha-
ride unit, GlcNAc (Figure 1B). This event produces a disaccha-
ride pentapeptide containing an undecaprenyl pyrophosphate
group. A major feat from the Walker Laboratory in 1999 showed
that Lipid I derivatives could be transformed with the glycosyl-
transferase MurG to form native Lipid II as well as a variety of de-
rivatives (Ha et al., 1999; Men et al., 1998). This in vitro enzymatic
conversion provided the first reliable method to produce work-
able quantities of Lipid II. Access to Lipid II, in turn, enabled
the mechanism of various glycopeptide antibiotics to be studied.
For example, vancomycin, as well as other glycopeptides such
as dalbavancin (Leimkuhler et al., 2005), has been shown to
inhibit TGs directly (Chen et al., 2003).
In 2017, a method to accumulate and isolate Lipid II directly
from cells was developed (Figure 2A) (Qiao et al., 2017). Qiao,
Kahne, and Walker developed a two-step extraction protocol of
Lipid II. Using a similar strategy as Strominger in 1958
and armed with the knowledge that PG TGs are responsible
for the last steps of PG biosynthesis (Figure 1B), antibiotics
that target these later steps were used to isolate native Lipid
II (Figure 2A). Cells treated with moenomycin or vancomycin,
which inhibit late-stage biosynthetic enzymes in the outer
leaflet of the plasma membrane (Chen et al., 2003; Kahne
et al., 2005; Taylor et al., 2006), accumulate a significant
amount of Lipid II (Figure 2A). However, when cells were
treated with a sublethal dose of fosfomycin, an antibiotic that
inhibits the first committed step in PG biosynthesis (Falagas
et al., 2016), Lipid II was undetectable (Qiao et al., 2017). The
ability to alter the amounts of PG biosynthetic intermediates
is a powerful tool for studying both upstream and downstream
effects of peptidoglycan synthesis, as evidenced by experi-
ments that have used the Lipid II isolation method to screen
for novel antibiotics, assess the function of critical PG biosyn-
thetic enzymes, and identify new enzymes involved in PG
biosynthesis (as discussed below).
To interrogate the production of Lipid II in a cellular context,
the Walker and Kahne labs have developed an easily accessible
technique to label Lipid II from bacterial cultures (Figure 2B)
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4. (Qiao et al., 2014). The method involves detection of Lipid II from
growing bacterial cells by treating the intermediate with peni-
cillin-binding protein 4 (PBP4) to allow for the installation of a
biotin tag. This low molecular weight PBP isolated from
S. aureus is a promiscuous transpeptidase that, in addition to
catalyzing bonds between adjacent muropeptides, can ex-
change the terminal D-Ala of the muropeptide with various D-
amino acids. In vitro, this technique can switch the terminal D-
Ala for biotinylated D-Lys (BDL); biotinylated Lipid II is then
detectable via western blot (Figure 2B). This technique provides
Figure 2. Biochemical and Chemical Biology Techniques for Intergorating Bacterial Peptidolgycans
(A) Two-step extraction method for isolating Lipid II from bacterial cultures.
(B) PBP4 transpeptidase mediated terminal D-Ala exchange with unnatural amino acids. Depicted is the incorporation of biotin-D-Lys (BDL) to the stem peptide of
Lipid II. Lipid II consists of a diphosphate (P) disaccharide backbone, GlcNAc (G), and MurNAc (M), with a pentapeptide chain: alanine (A), glutamate (E), and
lysine (K).
(C) Substituted cysteine accessibility method (SCAM) utilizes single cysteine mutations (orange circle) in a protein of interest in conjunction with two cysteine
reactive reagents: MTSES and NEM. MTSES cannot penetrate the membrane and will only react with periplasmic cysteines. NEM can penetrate the membrane
and react with both periplasmic and cytoplasmic cysteine residues. Using this method with a cysteine mutant library, it can provide topological information of a
protein, in this case a transmembrane protein.
(D) MurJ-pBPA photocrosslinking assay for detection of Lipid II/MurJ adducts. MurJ encoded with single pBPA mutations undergoes crosslinking with Lipid II
upon UV activation. After SDS-PAGE and electroblotting, crosslinked Lipid II is biotinylated in-gel via a BDL exchange reaction and then detected via blotting with
streptavidin-HRP.
(E) Subsequent lysis and a click reaction to attach a fluorophore allow for analysis of mycolate-protein interactions via metabolic incorporation of a bifunctional
TMM analog. N-x-AlkTMM-C15 is metabolically incorporated into the mycobacterial mycomembrane (MM). Covalent crosslinks with MM-associated proteins
are induced by UV activation. Subsequent lysis and a click reaction to attach a fluorophore allow for analysis by a variety of techniques.
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5. researchers with a streamlined process to obtain functionalized
Lipid II analogs that can be selectively detected, thus generating
a valuable assay for studying PG biosynthetic machinery and
screening antibiotics (Cochrane and Lohans, 2020).
In sum, there are at least three methods to access Lipid II and
its derivatives that have been reported: in vitro biochemical
methods using MurG, isolation of Lipid II from bacterial cultures,
and Lipid II biotinylation. Collectively, these methods have been
applied to diverse bacterial species: S. aureus, Bacillus subtilis,
Escherichia coli (Qiao et al., 2017), and M. smegmatis (Garcı́a-
Heredia et al., 2018) have all been successfully used for the Lipid
II isolation and/or visualization, despite variation in PG penta-
peptide chains (Qiao et al., 2017). Access to these substrates
will be important to fully understand the structural diversity pre-
sent in bacterial cell walls and to characterize pools of PG inter-
mediates during growth. These methods have been useful in
studying the mechanism of action of antibiotics, as mentioned
above with glycopeptides and more recently with lysobactin
(Lee et al., 2016). Research groups have also used them to study
the mechanisms of enzymes involved in PG biosynthesis. For
example, multiple labs have used the tools to biochemically
characterize the TG activity of the SEDS proteins, which have
been proposed as new potential targets for antibiotics (Cho
et al., 2016; Emami et al., 2017; Meeske et al., 2016; Rohs
et al., 2018; Sjodt et al., 2018; Taguchi et al., 2019b). The
methods of isolation and detection (Figures 2A and 2B) have
been used in combination in vitro to study the order of addition
of WTA precursors to PG intermediates (Figure 3B; this work
Figure 3. Critical Details for Understanding Peptidoglycan Isotopic Probe Incoroporation
(A) Cell wall depictions of S. aureus strains utilized to determine characteristic peaks of PG and TA.
(B) Assessing the ability of un-cross-linked and cross-linked PG as a substrate for WTA transfer by LcpB. Un-cross-linked PG oligomers featuring a pentaglycine
chain are bioenzymatically prepared with a SgtB mutant. The oligomers are then either modified with WTA via LcpB (top) or cross-linked with PBP4 (bottom).
Subsequently, these molecules are either subjected to transpeptidation with PBP4 or a ligation reaction with LcpB.
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6. will be discussed in detail in Radiolabeling). Finally, these
methods have been used to identify and study the mechanism
of Lipid II transport (Rubino et al., 2020) as discussed in detail
in the next section.
BIOCHEMICAL TOOLS TO INVESTIGATE THE PG
ENZYMES
For a long time, it was not understood how Lipid II translocates
across the lipid bilayer into the periplasmic space for incorpora-
tion into the PG meshwork. Multiple proteins were evoked to
perform this function, including the SEDS protein FtsW, which
was later shown to be a transglycosylase (Egan et al., 2020;
Ruiz, 2016; Young, 2014). MurJ was discovered by Ruiz and col-
leagues to be the elusive flippase (Ruiz, 2008; Sham et al., 2014).
In a series of elegant experiments utilizing substituted cysteine
accessibility method (SCAM) (Figure 2C) and the protein toxin
ColM (Touzé et al., 2012), Ruiz and collaborators were able to
create an assay that provided context-dependent monitoring
of Lipid II movement. Using an extensive single Cys mutant
MurJ library, the authors mapped the topology of MurJ, which
was then used in parallel with protein modeling to predict
possible dynamic conformations (Butler et al., 2013). Building
on these findings, Sham et al. used an in vivo assay with the addi-
tion of sulfonating reagents, ColM, or both in either wild-type
MurJ or the reversible MurJ mutant to formalize MurJ as the Lipid
II flippase (Sham et al., 2014).
With the knowledge that MurJ was the flippase, the biochem-
ical details of its activity were investigated using the Lipid II as-
says discussed above. MurJ proved challenging to study
because, as a flippase, it does not structurally alter the Lipid II
precursor when it transports. However, by using the Lipid II
detection method (Figure 2B), it was possible to monitor
changes in Lipid II’s movement. In 2018, Rubino et al. showed
that treatment with a protonophore to disperse the proton-
motive force caused Lipid II accumulation in E. coli, suggesting
that the transportation of Lipid II is coupled to an electrochemical
gradient. The effect of the protonophore mimicked controls
known to disrupt MurJ activity. Furthermore, they characterized
the conformation of MurJ when the membrane potential is dissi-
pated by probing individual cysteine residues in MurJ (Rubino
et al., 2018).
To determine what residues of MurJ are involved in transport,
photocrosslinking experiments were used to tether interacting
partners. Photocrosslinking has become an invaluable method
to determine protein-substrate interactions (Lancia et al., 2014;
Parker and Pratt, 2020; Wu and Kohler, 2019). Unnatural amino
acid incorporation can be used to install a photoactivatable
p-benzoyl-L-phenylalanine (pBPA) in the protein of interest—in
this case, MurJ. Excitation of pBPA using 350–365 nm light leads
to a reactive diradical that forms a C-H bond to any vicinal func-
tional group within a 3.1 Å reactivity radius (Lancia et al., 2014). In
2012, Okuda et al. utilized this method to determine specific sites
where LPS interacts with the LPS transport (Lpt) machinery
(Okuda et al., 2012). Briefly, they incorporated pBPA residues
in several Lpt transport proteins. After UV activation, photo-
crosslinked adducts were identified using immunoblotting with
LPS-specific antibodies. This method allowed them to take
chemical snapshots of LPS transport and determine that shut-
tling of LPS across the periplasm is accomplished through cyto-
plasmic ATP hydrolysis. The Ruiz group, in collaboration with the
Kahne lab, applied this methodology to probe MurJ flippase ac-
tivity (Figure 2D) (Rubino et al., 2020). They hypothesized that
pBPA incorporated into MurJ would prompt crosslinking to its
natural substrate Lipid II. However, the lack of antibodies for
Lipid II made it necessary to implement a method that allowed
for the detection of the crosslinked adduct. PBP4 was used to
incorporate a BDL after photocrosslinking Lipid II to MurJ-
pBPA mutants to allow for detection (Figure 2D). They applied
this protocol to investigate the role of three essential arginine
residues, located in the central cavity of MurJ, that were previ-
ously proposed to be key in recognizing the pyrophosphate of
Lipid II (Kuk et al., 2019). Using single and multiple Arg/Ala mu-
tants, they observed similar levels of crosslinking compared to
wild-type MurJ. However, these mutants displayed impaired
ability to flip Lipid II. This methodology permitted the observation
of the intermediate transport steps in living cells and provided
direct, biochemical evidence that the conserved arginine resi-
dues control Lipid II movement through MurJ. Thus, through
the combination of Lipid II chemical probes, genetic tools, and
biochemical conversions, the function of MurJ has been identi-
fied and the biochemical mechanisms of Lipid II transport are
rapidly being unveiled. This also highlights MurJ as an exciting
target for antibiotic development. The ability to track Lipid II in
cellular biochemistry assays was critical because it yielded
detailed biochemical information (i.e., protein residues, mem-
brane potential) that would not have been possible with other
methods.
METABOLIC INCORPORATION OF
PHOTOCROSSLINKING SUGARS
The proteins that bind the cell wall and its associated glycocon-
jugates are also potential antibiotic targets. In contrast to pro-
tein-mediated interactions, glycan recognition events are often
weak and short-lived. Additionally, glycans are not directly
genetically encoded and their biosynthesis is complex, so it is
challenging to use standard genetic engineering methods to
tag them. The incorporation of functionalized metabolites has al-
lowed a way to bypass these challenges (Campbell et al., 2007).
Several groups have introduced unnatural sugars containing
photoactivatable crosslinkers to the cell by hijacking carbohy-
drate metabolic pathways and capitalizing on enzyme promiscu-
ity (Tanaka and Kohler, 2008; Yu et al., 2012). The mycomem-
brane (Figure 1A, ‘‘MM’’) is attached to the mycobacterial cell
wall via AG and is a barrier to environmental, immune, and anti-
biotic insults. However, its protein composition has eluded
classic biochemical techniques for a long time, in part because
of the difficulty of cleanly separating the covalently bound myco-
membrane from other layers of the complex mycobacterial enve-
lope. Kavunja et al. recently developed the first photocrosslink-
ing probes for the mycomembrane to analyze mycolate-protein
interactions in vivo (Figure 2E) (Kavunja et al., 2020). They syn-
thesized a TMM analog that specifically incorporates into the
TDM portion of the mycomembrane via previously reported
conserved, substrate-promiscuous Ag85 mycoloyltransferases
(Fiolek et al., 2019). This analog contains a bifunctional linker
bearing a photoactivatable diazirine group and a clickable alkyne
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7. handle. After metabolic incorporation into the cell surface, myco-
bacteria were irradiated with UV light. The diazirine cross-linked
with neighboring proteins that were then enriched following click
ligation to an azide-fluorophore-biotin. This method allowed for
the identification of both known and previously undetectable my-
comembrane-resident proteins, as well as tracking them by in-
gel fluorescence. Similar techniques are likely to be useful in
future studies to interrogate different layers of the cell envelope,
including PG—especially as DeMeester and coworkers have
shown that PG precursors containing the diazirine cross-linkers
at the C2 position of MurNAc are accepted by the PG biosyn-
thetic enzymes (DeMeester et al., 2018). Previously, Sarkar
et al. exploited the MurF ligation process to insert a D-Ala- that
was biofunctionalized with an alkyne and photocrosslinking han-
dles into Lipid II to mine the protein-interacting partners (Sarkar
et al., 2016). Targeted acquisition of cell envelope interactomes
may reveal new potential targets for antibiotic therapies (Kavunja
et al., 2020).
SPECTROSCOPIC METHODS TO STUDY THE
CELL WALL
The stable isotopes 13
C and 15
N are effective as probes to eluci-
date structural features of the bacterial cell wall at the molecular
level using spectroscopic methods (Kim et al., 2014, 2015; Yang
et al., 2017). In this strategy, the macromolecules of biological in-
terest are unperturbed and uniform enrichment enhances signal
output of the NMR spectrum in a non-destructive manner (Ny-
gaard et al., 2015). Romaniuk and Cegelski utilized 13
C and
15
N cross-polarization magic angle spinning (CP/MAS) solid-
state NMR (ssNMR) to characterize the composition of uniformly
labeled PG and WTA in S. aureus (Romaniuk and Cegelski,
2018). This technique affords an alternative to solution-based
analytical methods that do not permit full characterization of
the highly insoluble material. Using this approach, spectra of pu-
rified PG and WTA isolates from wild-type and DtarO (a mutant
that is unable to synthesize WTA), respectively, were first used
to identify and quantify characteristic carbon peaks from each
component (Figure 3A). Since the sum of the two spectra repro-
duced the peak intensities of the intact cell wall sample, they
were able to quickly determine the composition ratio of TA to
PG. The masses for TA and PG calculated by this method
were consistent with those determined by phosphate analysis,
a more traditional but labor-intensive method. Subsequently,
the relative composition ratios of the two components in both
the stationary and the exponential phases were determined.
In the stationary phase, PG thickness increased while WTA
decreased. This finding was confirmed with selective labeling
using either D-[15
N]Ala (WTA) or [15
N]Gly (PG) in 15
N CP/MAS.
With these baselines in hand, they were able to validate that
this analytical method is amenable to determining the composi-
tion levels of TA and PG of cell walls with the antibiotic tunicamy-
cin, which inhibits WTA growth. Cegelski et al. have used similar
strategies with ssNMR to establish that vancomycin primarily
targets transglycosylation over transpeptidation using uniformly
13
C- and 15
N-labeled amino acids in S. aureus (Cegelski et al.,
2002). Changes to the D-alanine-pentaglycyl bridge-links were
unperturbed in the presence of vancomycin, suggesting that
transpeptidation is unaffected. Instead, this supports the idea
that vancomycin blocks transglycosylation and impedes translo-
cation of Lipid II into the periplasm. Rotational echo double reso-
nance (REDOR) and CP/MAS ssNMR have also been used to
discern perturbations to PG and WTA of S. aureus treated with
the cyclic decapeptide amphomycin, a drug that is effective
against superbugs such as multi-drug-resistant S. aureus and
vancomycin-resistant enterococci by targeting bactoprenol-
phosphate. Using REDOR, Singh et al. observed 15
N shifts in
bridge-links between Gly and L-Lys and the free side chain amine
of lysine, indicating that the compound induced PG thinning, the
accumulation of Park’s nucleotide, and a decrease in alanylation
of WTA (Singh et al., 2016). These data suggested that ampho-
mycin acts on the cell wall prior to transglycosylation. Overall,
CP/MAS and REDOR analysis of the bacterial cell wall permits
rapid assessment of whole-cell composition and can be applied
to monitor and determine the compositional perturbations
caused by antibiotics as showcased in the studies above.
In another example, Calabretta et al. employed 13
C radiola-
beled lipid-linked arabinofuranose donors to study Gram-inde-
terminate bacteria. These probes were biosynthetically incorpo-
rated into the arabinan layer of Corynebacterium glutamicum
and M. smegmatis strains unable to produce this polymer due
to genetic mutation or treatment with benzothiazinone antibi-
otics (Calabretta et al., 2019). This procedure circumvents the
requirement of metabolic processing of cell wall probes prior
to incorporation into the cell wall. Analysis of the soluble arabinan
isolated from these models retained characteristic peaks with 2D
NMR experiments (1
H-13
C HSQC, 1
H-13
C HMBC, and 1
H-13
C
HSQC-TOCSY). Though this study did not use intact cells to
gain structural details, it yielded a powerful tool that can be
applied to examine structural changes in the lesser-understood
mycobacterial cell envelope. Antibiotics are critically needed
against mycobacteria to combat public health threats such as
tuberculosis, which the Centers for Disease Control (CDC) has
recently identified as a top AMR threat (Bloom and Murray,
1992; CDC, 2017).
RADIOLABELING
Although NMR methods are useful in determining composition
of the cell wall, they do not inform on its biosynthesis. This can
be achieved using radioisotopes, as bacterial incorporation
of extremely sensitive ionizing radionuclides permits submicro-
molar (high fM to pM) detection of cell wall products and inter-
mediates of biosynthetic pathways in vivo. This sensitivity is
also useful in translational applications for disease diagnosis.
Here we discuss the use of 14
C and 11
C radionuclides in bac-
terial cell surface structures to discern mechanistic details of
PG synthesis and structure (long-lived) in addition to usage
as a diagnostic imaging tool of bacterial infection in vivo
(short-lived).
Long-Lived Radioisotopes
Elucidating some of the lesser-understood biosynthetic pro-
cesses has the potential to provide new targets for antimicro-
bials. TAs are important for the structure and virulence of
Gram-positive bacteria (Brown et al., 2013), but far less is known
about their synthesis compared to that of PG. Recently, Schaefer
et al. investigated the order and process by which WTA are
ll
1058 Cell Chemical Biology 27, August 20, 2020
Review

8. appended to PG. This is useful, as methicillin-resistant S. aureus
(MRSA) is resensitized to b-lactams when WTA biosynthesis is
inhibited (Campbell et al., 2011; Farha et al., 2013). They found
that WTA is ligated to un-crosslinked PG oligomers and that liga-
tion preference is due to steric restrictions of the LcpB WTA
ligase binding site (Schaefer et al., 2018). With isolated Lipid II
from S. aureus, featuring a pentaglycine chain for downstream
cross-linking, the group bioenzymatically prepared un-cross-
linked PG fragments (2–10 carbohydrates in length) with a trans-
glycosylase mutant (SgtBY181D
) that releases premature PG olig-
omers (Figure 3B). These fragments were then subjected to
transpeptidation with PBP4 followed by treatment with LcpB
(WTA ligase) and 14
C radiolabeled LIIA
WTA
, a truncated radiola-
beled WTA precursor, or just the latter to assess substrate pref-
erence. Detection by polyacrylamide gel electrophoresis (PAGE)
autoradiography showed that the cross-linked substrate is un-
able to be ligated to WTA, whereas the un-cross-linked PG frag-
ments serve as an acceptable substrate for the transfer of WTA
by LcpB. Additional experiments evaluating the structural re-
quirements for recognition co-crystalized the homolog TagT
from B. subtilis with either LIA
WTA
or LIIA
WTA
, which indicated a
putative binding groove narrow in size. The steric restrictions
of this site prompted Schaefer and coworkers to explore if the
stem peptide was a necessary feature of recognition by employ-
ing chitin, deacetylated chitin, and cellulose-based oligosaccha-
rides as probes of LcpB and TagT. Results indicated that the
stem peptide is not a necessary structural feature for recognition
of the transfer substrate; however, acetylation of the C2 position
of MurNAc is required. These radioisotopic probes enabled
Schaefer and coworkers to visually determine if PG intermedi-
ates were modified by WTA ligase in this enzymatic assay and
suggest the ligation order of WTA precursors to peptidoglycan
intermediates. With this fundamental information, new antibiotic
targets can be established as a means to stymie the ligation pro-
cess and ultimately disrupt the integrity of the cell wall of Gram-
positive bacteria.
Short-Lived Radioisotopes
Positron emission tomography (PET) radioisotopes such as 11
C,
13
N, 15
O, and 18
F are extremely sensitive tools that permit
whole-body imaging of physiological processes in deep-lying
tissues and organs with low nanogram quantities of probe. Un-
like stable isotopes, the radiation produced upon decay is
detectable outside of the human body. [18
F]-Fluorine deoxyglu-
cose (FDG) is a clinical PET probe frequently used to image cells
with higher energy requirements, such as oncogenic cells and
sites of inflammation (Zhuang and Alavi, 2002, Seminars in Nu-
clear Medicine, conference). [18
F]FDG accumulates because it
cannot be further metabolized after sequestration into the cell.
Since this probe is not bacteria-specific, improved imaging
technologies are required to detect active bacterial infections
in vivo (Ordonez and Jain, 2018, Seminars in Nuclear Medicine,
conference). To address this issue, Rosenberg, Ohliger, and
Wilson have developed clinically relevant 11
C radiotracers that
can distinguish between sterile inflammation and active infec-
tion by metabolically incorporating into both clinically relevant
pathogenic Gram-positive and Gram-negative bacteria (Neu-
mann et al., 2017; Parker et al., 2020; Stewart et al., 2020), uti-
lizing D-amino acid probe incorporation (de Pedro et al., 1997).
High incorporation of D-[14
C]Met in both S. aureus and E. coli
in the stem peptide of PG led to the development of a 11
C-syn-
thetic probe from a D-homocysteinethiolactone precursor
(Figure 4). Earlier experimentation had shown that exogenous
D-Met is amply integrated into the PG polymer of stationary bac-
teria (Caparrós et al., 1992; Lam et al., 2009). Moreover, a robust
synthetic strategy and efficient labeling probe for D-Ala and
D-Glu were not readily available. Using the L-[11
C]-Met tracer
for imaging in mice modeling myositis, they were able to detect
and differentiate between bacterial infection and sterile inflam-
mation, modeled by heat-killed bacteria, in the deltoid muscle
with great sensitivity. These probes are likely to be especially
useful for regions of the body that are normally sterile such as
the musculoskeletal, biliary, and nervous systems, as there will
not be significant background from the commensals. The syn-
thetic efficiency of the probe was later improved for use in the
clinical setting by using automated synthesis (Neumann et al.,
2017; Parker et al., 2020; Stewart et al., 2020). Most recently,
Parker et al. expanded the tracers to include amino acids
canonically incorporated into the stem peptide, D-[3-11
C]-Ala
and D-[3-11
C]-Ala-D-Ala probes (Figure 4). In vivo studies utilized
the D-Ala probe, since D-Ala showed anywhere from 2-3 times
greater uptake in E. coli and S. aureus over the dipeptide. More-
over, it was determined that the D-Ala probe generated high
levels of incorporation with the previous panel of bacteria tested
in the 2019 paper and that these probes were highly selective for
bacterial cells over mammalian cells. Infectious imaging chal-
lenge experiments also confirmed the ability of the D-Ala probe
to distinguish between live bacterial infection and sterile
Figure 4. Radiolabels in Living Bacteria’s Peptioglycans
Incorporation of radiolabeled probes, D-[methyl-11
C]-Met or D-[3-11
C]-Ala, post-biosynthetically via transpeptidases in clinically relevant bacteria to assess use
for in vivo labeling of active bacterial infections.
ll
Cell Chemical Biology 27, August 20, 2020 1059
Review

9. inflammation in mouse models. The tracer was able to monitor
cell wall-acting antibiotic (ampicillin) treatment of an E. coli bac-
terial infection, image infection in the intervertebral space, and
proficiently screen for pneumonia in mice. Thus, this class of
radiolabeled probes yielded an indispensable bacteria-specific
imaging strategy for pathogenic bacteria in difficult-to-access
physiological niches in clinical settings compared to currently
approved methods.
CONCLUSION
Biochemists have been using small molecules to study bacterial
cell walls since the 1800s. As improved probes have become
available, the ability to study the biochemical phenomena has
exploded. Here we have highlighted examples of chemical
probes that were used to reveal fundamental cell wall biochem-
istry or, in some cases, to identify bacterial pathology in a living
host. There is a rich intersection between chemical biology and
cell walls. It is this intersection that will be powerful in tackling
the problem of antibiotic resistance, as a means to identify
new targets and to characterize the mechanism of action for
promising compounds. We highlighted the recent advances in
the field such as monitoring PG precursors to elucidate MurJ,
TG, and TP activity; establishing new protein targets for anti-
biotic development through photocrosslinking techniques;
applying NMR spectroscopy to capture native cell wall compo-
sition; and demonstrating a highly sensitive method of detection
of cellular processes using radiolabels in vitro and in vivo. In
conjunction with current visualization and imaging techniques,
chemical, biochemical, and hybrid methods have the potential
to provide both mechanistic and structural information for valu-
able targets.
ACKNOWLEDGMENTS
This project was supported by a grant from the Glycoscience Common Fund
(U01 CA221230) and the Delaware COBRE program, as well as a grant from
the National Institute of General Medical Sciences-NIGMS (P20GM104316).
C.L.G. is a Pew Biomedical Scholar, Sloan Scholar, and Camille Dreyfus
Scholar and thanks the Pew Foundation, the Sloan Foundation for Science
Advancement, and the Dreyfus Foundation for support. M.S.S. thanks NIH
R21 AI144748 and NIH DP2 AI138238 for financial support. S.H. thanks the
NIH Chemistry-Biology Interface Program (GM133395) and the University of
Delaware Dissertation Fellowship for funding support.
DECLARATION OF INTERESTS
C.L.G. and M.S.S. have pending patents and patents relevant to this work.
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