of 12
Cohesive
Living
Bacterial
Films
with Tunable
Mechanical
Properties
from
Cell Surface
Protein
Display
Published
as part of ACS Synthetic
Biology
special
issue “Materials
Design
by Synthetic
Biology”.
Hanwei
Liu,
#
Priya
K. Chittur,
#
Julia A. Kornfield,
and David
A. Tirrell
*
Cite This:
ACS Synth.
Biol.
2024, 13, 3686−3697
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*
Supporting
Information
ABSTRACT:
Engineered
living materials
(ELMs)
constitute
a novel class of functional
materials
that contain
living organisms.
The
mechanical
properties
of many such systems
are dominated
by the polymeric
matrices
used to encapsulate
the cellular
components
of the material,
making
it hard to tune the mechanical
behavior
through
genetic
manipulation.
To address
this issue, we have
developed
living materials
in which mechanical
properties
are controlled
by the cell-surface
display of engineered
proteins.
Here, we
show that engineered
Esherichia
coli
cells outfitted
with surface-displayed
elastin-like
proteins
(ELPs, designated
E6) grow into soft,
cohesive
bacterial
films with biaxial moduli
around
14 kPa. When subjected
to bulge-testing,
such films yielded
at strains of
approximately
10%. Introduction
of a single cysteine
residue near the exposed
N-terminus
of the ELP (to afford a protein
designated
CE6) increases
the film modulus
3-fold to 44 kPa and eliminates
the yielding
behavior.
When subjected
to oscillatory
stress, films
prepared
from
E. coli
strains bearing
CE6 exhibit modest
hysteresis
and full strain recovery;
in E6 films much more significant
hysteresis
and substantial
plastic deformation
are observed.
CE6 films heal autonomously
after damage,
with the biaxial modulus
fully restored
after a few hours. This work establishes
an approach
to living materials
with genetically
programmable
mechanical
properties
and a capacity
for self-healing.
Such materials
may find application
in biomanufacturing,
biosensing,
and bioremediation.
KEYWORDS:
engineered
living
materials,
protein
surface
display,
tunable
mechanical
properties,
self-healing
materials,
disulfide
engineering
INTRODUCTION
In nature,
microorganisms
including
bacteria
can autono-
mously
assemble
into hierarchical
biofilms,
1
composed
in part
of living cells that can sense environmental
stress
2
and catalyze
reactions,
3,4
and in part of extracellular
polymeric
matrices
(EPS)
5
that are secreted
by cells and composed
of proteins,
lipids, polysaccharides,
and nucleic
acids.
6
These polymeric
matrices
create microenvironments
that help bacteria
survive
environmental
challenges.
7
Inspired
by natural
biofilms
and
other biomaterials,
the field of engineered
living materials
lies
at the interface
between
materials
science
and synthetic
biology.
Through
manipulation
of genetic
information,
organisms
can be directed
to assemble
into materials
that
possess
desirable
characteristics
of living systems,
such as
autonomous
assembly,
adaptiveness
to environmental
stimuli,
and self-healing.
8
12
Recently,
several
biofilm-inspired
living
materials
have been reported,
in which researchers
either
rewired
the production
of natural
biopolymers
like curli
fibrils
13,14
or cellulose,
15,16
or encapsulated
bacteria
in
synthetic
polymeric
matrices.
17,18
Proteins
and peptides
have
been fused to curli protein
monomers
and assembled
into
materials
that exhibit novel catalytic
functions.
19,20
Cellulose-
Received:
August
2, 2024
Revised:
October
22, 2024
Accepted:
October
23, 2024
Published:
November
1,
2024
Research
Article
pubs.acs.org/synthbio
© 2024 The Authors.
Published
by
American
Chemical
Society
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producing
bacteria
assembled
at the air
water
interface
21,22
form pellicles
that can be collected
and processed
via 3D
printing.
15
Encapsulating
bacteria
in synthetic
polymer
net-
works can protect
the cells from environmental
insults while
preserving
sensitivity
to environmental
stimuli.
18
In such
materials,
the polymeric
matrix
dominates
the mechanical
properties,
while the encapsulated
cells serve as environmental
sensors.
Matrix-free
living materials
have also been described.
In such
systems,
bacteria
are engineered
to express
associative
proteins
and display
them on the cell surface,
forming
cell aggregates.
This approach
requires
neither
matrix secretion
nor synthetic
polymers,
so that living materials,
in principle,
can be
generated
from a single cell. Materials
of this kind have been
developed
on the basis of nanobody-antigen
interactions
in
E.
coli
,
23,24
and through
engineering
Caulobacter
crescentus
by
modifying
its RsaA protein.
25,26
In prior examples,
cells were
grown in liquid culture and then collected
and assembled
into
cm-scale
structures.
However,
the growth
of centimeter-scale
living cohesive
bacterial
films with tunable
mechanical
properties
on solid surfaces,
utilizing
surface-displayed
self-
associative
proteins,
has not been reported.
Here, we explore
the possibility
of
growing
a cohesive
bacterial
film directly
on a solid substrate
using a previously
developed
protein
surface-display
system
27
to engineer
E. coli
to display elastin-like
proteins,
with the introduction
of a single
amino acid that enables
covalent
links between
neighboring
cells. Surface
display of unstructured
elastin-like
proteins,
150
amino acids in length, enabled
bacterial
cells to form cohesive
films. The introduction
of a single cysteine
residue
near the
Figure
1.
Cohesive
living films made from engineered
bacteria.
(A) Schematic
representations
of engineered
bacteria
and types of intercellular
adhesion.
(B) Suction-coating
method
for preparation
of bacterial
films. (1) Overnight
planktonic
culture of bacteria
of interest
was pipetted
onto a
perforated
polycarbonate
filter with a 0.2
μ
m diameter
pore size. (2) Vacuum
filtration
was applied
to remove
liquid media and retain bacteria
on
the filter. (3) The filter with bacterial
coating
was then transferred
to a fresh LB agar plate daily for 7 days. An image of a 7-day-old
CE6-AT
bacterial
film on a polycarbonate
filter on an LB agar plate. Scale bar: 1 cm. (C) Microscopy
image of a microtomed
cross-section
of a 7-day-old
CE6-AT
film that expresses
mWasabi
as a fluorescence
marker.
Scale bar: 10
μ
m. (D) Microscopy
image of a 7-day-old
CE6-AT
film microtome
cross-section
immunostained
by an anti-His-tag
antibody
conjugated
to Dylight-488.
Scale bar, 10
μ
m. The image shown in the upper left corner is
an enlargement
of the dashed-square
portion
of the image in the lower left corner.
ACS Synthetic
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Research
Article
https://doi.org/10.1021/acssynbio.4c00528
ACS Synth.
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2024, 13, 3686
3697
3687
exposed
N-terminus
of the displayed
elastin improved
the
mechanical
performance
of the films, presumably
by forming
intercellular
disulfide
bonds. The resulting
films without
the
cysteine
residue
yield when nominal
stress exceeds
1 to 2 kPa;
in contrast,
films with the cysteine
did not undergo
yielding
even when subjected
to nominal
stress of 4 to 5 kPa, survived
multiple
loading
cycles without
permanent
deformation,
and
exhibited
self-healing
after fracture,
with some recovering
their
moduli
fully within a few hours.
RESULTS
AND
DISCUSSION
Design
of Surface-Displayed
Proteins.
We expressed
proteins
on the
E. coli
surface
using an autodisplay
system
previously
used to display a wide variety of proteins,
including
enzymes
and vaccine
epitopes,
28
31
as well as proteins
that
drive the formation
of bacterial
aggregates
in planktonic
culture.
27
In these cellular
constructs,
the protein
to be
displayed
is inserted
between
a PelB secretion
peptide
and the
autotransporter
(AT) outer membrane
protein
(Figure
S1).
Both of the displayed
proteins
employed
in this work carry
6xHis tags and elastin-like
(E) peptide
domains
previously
used to prepare
protein-based
hydrogels
32
34
and to engineer
microbial
assembly
in planktonic
culture.
27
Because
we used
six copies of a 25-amino
acid E domain,
which had proven
effective
in our earlier work in planktonic
systems,
we refer to
this construct
as E6-AT
(Figure
1A, center).
The CE6-AT
construct
differs from E6-AT
by a single amino
acid
a
cysteine
(C) residue
placed between
the 6xHis tag and the E6
domain
(Figure
1A, right). The goal of introducing
cysteine
near the N-terminus
of the displayed
protein
was to enable the
formation
of intercellular
covalent
disulfide
bonds.
These
surface-displayed
proteins
(sequences
given in Table S2) were
encoded
into a pQE80
plasmid
backbone
under the control
of
a T5-lac promoter.
In the
E. coli
DH10B
cells used in this
work, the T5-lac promoter
drives constitutive
expression
of the
surface-displayed
protein;
no inducer
is required.
Protocol
for Growing
Bacterial
Films.
A 200-
μ
L
volume
containing
approximately
10
8
cells grown from a single colony
on an agar plate was pipetted
onto a 2.5 cm diameter
polycarbonate
membrane
filter with 0.2
μ
m pore size, covering
a circular
area of roughly
90 mm
2
(Figure
1B, left). Vacuum
filtration
was applied
to remove
the liquid medium
and retain
the bacterial
cells on the membrane
filter. The filter was
transferred
to an LB agar plate, which provides
nutrition
for
bacterial
growth
and antibiotics
to maintain
the expression
of
the plasmid.
Nutrition
was replenished
by transferring
the filter
to a fresh LB agar plate every day. Bacterial
films were
characterized
after 7 days of growth.
Structure
of Bacterial
Films.
To probe the structure
of
bacterial
films, we introduced
a second
plasmid
to drive the
constitutive
expression
of a fluorescent
protein.
An image of a
microtomed
cross-section
of a CE6-AT
film that expresses
mWasabi
shows that 7 days of growth
(described
above)
provides
films approximately
70
μ
m thick that contain
densely
Figure
2.
Mechanical
properties
of engineered
bacterial
films measured
by the ramp bulge test. (A) Schematic
of the bulge test device, fabricated
as
two parts that are separated
to load a sample in the chamber
at the center of the device; the top and bottom
parts seal with vacuum
grease. When
loaded and sealed, “port to reservoir
1” connects
to a reservoir
of fluid that is used to control
pressure
on the top face of the sample (not shown),
and “port to reservoir
2” permits
control of the pressure
on the bottom
face of the sample (Figure
S5). Gray layers are acrylic; blue indicates
etched
channels
(light blue channel
to the top face of the sample;
darker blue channel
to the bottom
face of the sample);
green shaded
area indicates
coverslip
glass; layers are bonded
using epoxy. Horizontal
channels
are longer than shown. (B) Schematic
of loading
the bacterial
film sample into
the bulge test device and imaging
of the deformed
film by optical coherence
tomography
(OCT).
The punched
bacterial
film sample
(3 mm
diameter)
supported
by two washer-shaped
disks was transferred
to the central chamber
of the device. A thin O-ring sealed this “sandwich”
to the
top half of the device. When pressure
in reservoir
2 was larger than that in reservoir
1, a net pressure
upward
was applied
to the bacterial
film; the
film bulged upward
and was imaged
by OCT. (C) Three representative
cross-sectional
OCT images (right, numbered)
acquired
during the bulge
test of a CE6AT
film under linearly
increasing
pressure,
at the points indicated
on the stress
strain
curve (Right).
Numbers
with stars indicate
data
points corresponding
to the images.
Scale bar 500
μ
m. (D) Stress versus strain curves of CE6-AT
and E6-AT films. Number
of curves shown for
CE6-AT
films: 3. Number
of curves shown for E6-AT films: 3. (E) Biaxial modulus
values of CE6-AT
and E6-AT films. Number
of replicates
for
CE6-AT:
7. Number
of replicates
for E6-AT films: 4. Error bars represent
1 standard
deviation.
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packed bacterial
cells (Figure
1C). To ensure that the elastin-
like proteins
were in fact displayed
at the bacterial
surface
in
these films, we stained
microtomed
sections
with the anti-
6xHis antibody
conjugated
to Dylight-488.
Staining
by
fluorescent
antibodies
is evident
across the full thickness,
indicating
the expression
of CE6-AT
protein
throughout
the
film (Figure
1D). At higher magnification
(Figure
1D, lower
left), we see evidence
of antibody
staining
surrounding
the cell
body and appearing
to form a ring when the cell is viewed on-
axis. These images suggest
that His-tagged
protein
is expressed
at the cell surface,
consistent
with the expected
surface display
of CE6-AT.
Similar cross-sectional
imaging
was applied
to E6-
AT films, and flow cytometry
was used to analyze
cell
suspensions
derived
from empty-vector
controls
and E6-AT
films (Figures
S2 and S3). Empty-vector
control
films were too
fragile to allow microtome-sectioning,
staining,
and imaging,
but flow cytometry
confirms
that antibody-staining
of E6-AT
cells is roughly
2 orders of magnitude
stronger
than that of
controls.
Taken together,
these results confirm
that E6-AT and
CE6-AT
films consist of densely
packed
cells with elastin-like
proteins
displayed
at the cell surface and expressed
throughout
the depth of the film.
Erosion
Assay
of Film Cohesion.
As a first test of film
cohesion,
we applied
a simple erosion
assay. Seven-day-old
bacterial
films were immersed
in 7 mL of PBS buffer in six-well
plates and placed on a rocking
platform
using a 15
°
tilt angle at
15 turns per min. Optical
densities
(OD
600
) of buffers
in
contact
with bacterial
films were measured
at 1 and 24 h. After
1 h of erosion
in PBS, control
films without
surface
protein
expression
reached
an OD
600
value above 0.1, in contrast
to
OD
600
< 0.001 for both E6-AT and CE6-AT.
After 24 h of
erosion,
the supernatant
OD
600
reached
values greater than 0.3
for the control,
and no film was visible. In contrast,
E6-AT and
CE6-AT
films remained
intact, and the supernatant
OD
600
remained
below 0.01 (Figure
S4). These results confirmed
the
role of surface-displayed
proteins
in bacterial
film cohesion.
Determination
of Small-Strain
Biaxial
Modulus.
To
characterize
the mechanical
properties
of the cohesive
bacterial
films, we constructed
a custom
millifluidic
device suitable
for
applying
a “bulge test” to the films (Figure
2A). The device
imposed
hydrostatic
pressure
differences
in the Pa
kPa
range
across thin, freely suspended
film samples,
and was equipped
with an optical
coherence
tomography
(OCT)
system
to
quantify
changes
in film shape, which were analyzed
to extract
mechanical
properties.
35
A 3-mm diameter
sample,
cut from a
7-day-old
bacterial
film, was gently covered
with a drop of
buffer to relieve capillary
forces, lifted on a 3 mm-diameter
transmission
electron
microscopy
(TEM)
disk having a 1.5
mm diameter
aperture,
and placed into the central chamber
of
the bulge test device, where another
3 mm disk with a 1.5 mm
diameter
aperture
was placed on it (Figure
2B). Channels
in
the device allowed
for the independent
modulation
of the
hydrostatic
pressure
applied
to the top and bottom
faces of the
film, via changes
in fluid levels in two external
reservoirs,
“Reservoir
1” and “Reservoir
2” (Figures
2A and S5), with
associated
hydrostatic
pressures
p
1
and
p
2
. Thus, when the
liquid level in “Reservoir
2” was higher than that in “Reservoir
1” (
p
2
>
p
1
), the sample bulged upward
(Figures
2B and S5).
To increase
the pressure
difference
p
2
p
1
at a constant
rate, a syringe pump was used to add liquid to Reservoir
2 at a
constant
rate, imposing
a “ramp” pressure
time
profile. In the
present
experiments,
PBS buffer or HEPES
buffer was added at
Figure
3.
Disulfide
bonds play an essential
role in enhancing
the mechanical
properties
of CE6-AT
films. (A) Schematic
of TCEP reduction
of
disulfide
bonds and dye-labeling
of free thiols. (B) Absorption
intensity
after dye-labeling
of CE6-AT
and E6-AT films with or without
treatment
with TCEP. Number
of replicates
for each group: 3. Error bars represent
one standard
deviation.
(C) Biaxial modulus
values for E6-AT and CE6-
AT films with or without
treatment
with TCEP. Number
of replicates
for E6-AT TCEP+
= 4; for all other groups
n
= 3. Error bars represent
one
standard
deviation.
(D) Stress versus strain curves of E6-AT and CE6 AT films with or without
treatment
with TCEP. Number
of curves shown for
each group: 2.
**
represents
statistically
significant
differences
in mean values with
p
< 0.01. ns indicates
that differences
in mean values are not
statistically
significant.
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a flow rate of 70 mL/min
(with 60 mL syringes
used as
reservoirs,
the resulting
rate of pressure
increase
was 20.4 Pa/s;
see Figure S5 for additional
details)
until the total volume
transferred
reached
55 mL. The flow rate was chosen to be the
maximum
accommodated
by the syringe
pump to minimize
perfusion
through
the biofilm
for the duration
of the
experiment.
Simultaneously,
OCT images
were collected
(with an acquisition
time of 19 ms/image).
The images were
subsequently
processed
using a custom
script to extract the
evolution
of film stress and strain during the test (Figure
4C,
detailed
in the Materials
and Methods
section
and Supporting
Information).
Three stress
strain
curves for CE6-AT
films and three
curves for E6-AT films (each curve representing
a biological
replicate)
show an approximately
linear relationship
between
stress and strain for CE6-AT
films and a distinct
yielding
behavior
for E6-AT.
All E6-AT films were tested until failure,
while none of the CE6-AT
films failed even at a maximum
applied
pressure
of 960 Pa. We also examined
empty-vector
control
films, which were not engineered
for protein
surface
display;
those films were not cohesive
and could not be loaded
into the test apparatus.
Biaxial moduli were estimated
from the
initial linear portion
of each curve (below 5% strain) and were
higher for CE6 than for E6, as evident
in the steeper
slopes of
their stress
strain
curves. The average
biaxial modulus
of CE6-
AT films was determined
to be 44.0
±
5.6 kPa, approximately
three times that of E6-AT films (14
±
2.1 kPa). E6-AT films
also typically
yielded
and transitioned
to a plastic response,
eventually
failing before the end of the test, while CE6 films
showed
no evidence
of yielding
within the pressure
regimes
that we were able to test. The differences
in the yielding
behavior
of E6-AT films and CE6-AT
films were also observed
when we tried to peel films from their membrane
supports
using tweezers
without
providing
an aqueous
medium
to wet
the newly exposed
surfaces;
E6-AT
films were observed
to
stretch and break, while CE6-AT
films could be peeled from
the filter intact. Observation
of CE6 failure was rare, and in our
hands only occurred
once (for a film that was accidentally
left
in 1
×
PBS buffer for 4 h, Figure S6). Unlike the E6 films that
yielded,
the one CE6 film that failed exhibited
brittle failure
(Figure
S6). Videos
of OCT-scanned
“Ramp”
bulge tests of
E6-AT and CE6-AT
films, peeling
experiments,
and the only
observed
failure event of a CE6-AT
film are attached
in Videos
S1
S5;
video frame rates are rates of actual acquisition.
Mechanical
Strength
of CE6-AT
Films
Can
Be
Reduced
Chemically.
Relative
to E6-AT
films, CE6-AT
films were stronger
and stiffer (Figure
2D,E), supporting
our
hypothesis
that covalent
disulfide
bonds between
cells enhance
the mechanical
properties.
To assess the potential
role of
factors other than intercellular
covalent
bonds in enhancing
the
mechanical
properties
of CE6-AT
films, the water content
(Figure
S7), viability
of bacterial
cells (colony-forming
units
per unit mass of film, Figure S8), and protein
expression
levels
(Figures
S10) were all measured.
In each case, we found
similar results for the E6-AT and CE6-AT
films.
Disulfide
bonds can be cleaved
with a variety of reducing
reagents.
We chose odorless
tris(2-carboxyethyl)phosphine
(TCEP)
to reduce and count disulfide
bonds in bacterial
films.
Free thiols before and after reduction
can be capped
with a
maleimide
dye (Figure
3A). If CE6-AT
forms disulfide
bonds,
then CE6-AT
films treated
with TCEP should exhibit higher
maleimide-dye
labeling
than either CE6-AT
without
TCEP
treatment
or E6-AT films irrespective
of TCEP treatment.
We
used 50 mM TCEP in 20 mM HEPES
pH 7.0 buffer to treat
preweighed
films for 1 h (TCEP+
samples).
Control
films were
not treated with TCEP but were immersed
in 20 mM HEPES
buffer at pH 7.0 for 1 h (TCEP-
samples).
The respective
buffers were then removed,
and films were treated with 50
μ
M
Dylight
633-maleimide
in HEPES
buffer at pH 7.0 for 30 min.
Films were rinsed three times with 500
μ
L of HEPES
buffer
and then lysed. The absorbance
of the lysate at 633 nm was
measured
and normalized
by the wet mass of the treated film.
CE6-AT
films treated
with TCEP
exhibited
the strongest
labeling
(Figure
3B), suggesting
that these films contain
the
largest concentration
of free thiols (while CE6-AT
and E6-AT
films had similar Dylight-maleimide
absorbance
in the absence
of TCEP
treatment).
This result suggests
that surface-
displayed
thiols in CE6-AT
films are predominantly
in the
oxidized
(disulfide)
form. We note that this procedure
reveals
total disulfide
bonds; the percentage
of intercellular
disulfide
bonds cannot be determined
by this method.
We estimated
the number
of CE6-AT
proteins
per cell by
assuming
that the difference
in labeling
intensity
between
the
CE6 TCEP+
group and the E6 TCEP+
group represents
reduced
CE6-AT
proteins.
The calculation
method,
discussed
in Supporting
Note 4 and Figure S9, yields an estimate
of 2.5
×
10
5
CE6-AT
proteins
per cell. This result agrees with those of
quantitative
Western
blotting
(Supporting
Note 5 and Figure
S11), which also gives values of around
2.5
×
10
5
E6-AT and
CE6-AT
proteins
per cell. Assuming
the surface area of a single
E.
coli
cell is 6
μ
m
2
36
the density
of surface-displayed
engineered
protein
is approximately
4
×
10
4
proteins
per
μ
m
2
. Ajo-Franklin
and coworkers
recently
reported
a
C.
crescentus
-based
living material
25
with RsaA fused to an elastin-
like-peptide
displayed
at a density
of 1.4
×
10
5
proteins
per
μ
m
2
, which yielded
concentrated
cell suspensions
with shear
storage moduli of
14 kPa.
37
39
The display density estimated
here for E6-AT
and CE6-AT
films is consistent
with these
results,
as is the modulus
we’ve measured
for CE6-AT.
For a
given material,
the biaxial modulus
is greater
than Young’s
modulus,
which is typically
three times the shear modulus,
making
the 44 kPa biaxial modulus
of CE6-AT
comparable
to
the 14 kPa shear modulus
reported
by Ajo-Franklin
and
coworkers.
Next, we addressed
the question
of whether
disulfide
bonds
are responsible
for the enhanced
mechanical
properties
of the
CE6-AT
films. We treated
CE6-AT
and E6-AT films for 30
min either with 50 mM TCEP in 20 mM HEPES
buffer at pH
7.0 (TCEP+)
or with 20 mM HEPES
buffer at pH 7.0 (TCEP-
control).
Ramp bulge tests were then performed
on both the
TCEP+
and TCEP- groups in 20 mM HEPES
buffer at pH 7.0.
TCEP appeared
to have a minimal
effect on the mechanical
properties
of E6-AT
films, while CE6-AT
films were very
sensitive
to TCEP treatment.
As shown in Figure 3C, after the
treatment
of CE6-AT
films with 50 mM TCEP for 30 min (a 1
h treatment
was attempted,
but it made CE6-AT
too fragile to
load into the bulge test apparatus),
the biaxial
modulus
dropped
from above 70 kPa to just slightly
above 40 kPa.
Interestingly,
even after TCEP treatment,
CE6-AT
films still
showed
no evidence
of yielding
or failure under the conditions
used for bulge testing (Figure
3D) and exhibited
biaxial moduli
larger than those of E6-AT films (Figure
3C). These results
suggest
that the TCEP reduction
is incomplete
after 30 min.
OCT imaging
revealed
no structural
changes
in reduced
films.
Viscoelastic
Behavior
of Bacterial
Films.
We inves-
tigated the elasticity
and changes
in properties
over multiple
ACS Synthetic
Biology
pubs.acs.org/synthbio
Research
Article
https://doi.org/10.1021/acssynbio.4c00528
ACS Synth.
Biol.
2024, 13, 3686
3697
3690