Acoustic Biosensors for Ultrasound Imaging of Enzyme Activity

Anupama Lakshmanan

1,#

Zhiyang Jin

2,#

Suchita P. Nety

Daniel P. Sawyer

Audrey Lee-

Gosselin

Dina Malounda

Margaret B. Swift

David Maresca

Mikhail G. Shapiro

1,*

Division of Chemistry and Chemical Engineering, California Institute of Technology, Pasadena,

CA-91125, USA

Division of Engineering and Applied Sciences, California Institute of Technology, Pasadena,

CA-91125, USA

Division of Biology and Biological Engineering, California Institute of Technology, Pasadena,

CA-91125, USA

Abstract

Visualizing biomolecular and cellular processes inside intact living organisms is a major goal of

chemical biology. However, existing molecular biosensors, based primarily on fluorescent

emission, have limited utility in this context due to the scattering of light by tissue. In contrast,

ultrasound can easily image deep tissue with high spatiotemporal resolution, but lacks the

biosensors needed to connect its contrast to the activity of specific biomolecules such as enzymes.

To overcome this limitation, we introduce the first genetically encodable acoustic biosensors -

molecules that ‘light up’ in ultrasound imaging in response to protease activity. These biosensors

are based on a unique class of air-filled protein nanostructures called gas vesicles, which we

engineered to produce non-linear ultrasound signals in response to the activity of three different

protease enzymes. We demonstrate the ability of these biosensors to be imaged

in vitro

, inside

engineered probiotic bacteria, and

in vivo

in the mouse gastrointestinal tract.

INTRODUCTION

Virtually every biological process in living organisms involves dynamic changes in the

concentration or activity of specific molecules. Visualizing these changes within the context

of intact living tissues is critical to expanding our understanding of biological function and

developing next-generation medicines. A large repertoire of genetically encoded fluorescent

sensors has been developed to image specific molecular and cellular events

–

. However,

deploying such biosensors in living organisms is challenging due to the limited penetration

Correspondence should be addressed to MGS: mikhail@caltech.edu, Phone: 626-395-8588 or 617-835-0878, 1200 E. California

Blvd, MC 210-41, Pasadena, CA 91125.

Contributed equally

AUTHOR CONTRIBUTIONS

A.L. and M.G.S conceived the study. A.L., Z.J. and S.P.N. designed and planned experiments. A.L., Z.J., S.P.N., D.P.S., A.L-G.,

M.B.S. and D. Mal. conducted the experiments. Z.J., D.P.S. and D. Mar. wrote the MATLAB scripts for ultrasound imaging and data

processing. A.L, Z.J. and M.G.S. analyzed the data. A.L., Z.J. and M.G.S wrote the manuscript with input from all authors. All authors

have given approval to the final version of the manuscript.

COMPETING FINANCIAL INTERESTS

The authors declare no competing financial interests.

HHS Public Access

Author manuscript

Nat Chem Biol

. Author manuscript; available in PMC 2021 March 01.

Published in final edited form as:

Nat Chem Biol

. 2020 September ; 16(9): 988–996. doi:10.1038/s41589-020-0591-0.

Author Manuscript

of light in tissue

. In contrast, non-invasive techniques such as ultrasound are capable of

imaging deep tissues with high spatial and temporal resolution (below 100 μm and 1 ms,

respectively)

. However, ultrasound currently lacks the sensors needed to observe dynamic

molecular activity.

Here, we introduce molecular biosensors for ultrasound based on gas vesicles (GVs), a

unique class of air-filled protein nanostructures that were recently established as genetically

encodable imaging agents for ultrasound

. GVs evolved in certain aquatic microbes as a

means to regulate cellular buoyancy for optimal photosynthetic illumination

. GV

nanostructures comprise a 2 nm-thick protein shell enclosing an air-filled compartment, with

genetically determined widths between 45–250 nm and lengths of several hundred nm

The low density and high compressibility of GVs relative to surrounding aqueous media

allows these proteins to scatter sound waves and thereby produce ultrasound contrast when

injected into the body or expressed heterologously in engineered cells

We hypothesized that we could engineer GV-based biosensors that dynamically change their

ultrasound contrast in response to the activity of specific biomolecules. This possibility

arises from the recent discovery that GVs’ acoustic properties can be modified at the level of

their constituent proteins

. In particular, the scaffolding protein GvpC, which sits on the

GV surface (Fig. 1a) and provides structural reinforcement

, can be modified at the level of

its amino acid sequence to change GV mechanics. For example, shortening or removing

GvpC makes GVs less rigid, allowing them to buckle more easily under acoustic

pressure

. This reversible buckling produces nonlinear ultrasound contrast, which

appropriate ultrasound pulse sequences readily distinguish from the linear signals produced

by non-buckling GVs and background tissue

As an initial target for acoustic biosensor development, we chose proteases - an important

class of enzymes involved in many aspects of cellular signaling, homeostasis, disease,

therapy and synthetic biology

–

. While these enzymes were the targets of some of the

first fluorescent biosensors

, and continue to be a major focus of sensor engineering

no acoustic biosensors of protease activity have been developed. We postulated that by

engineering variants of GvpC incorporating amino acid sequences that are recognized and

acted upon by specific proteases, we could generate GVs whose nonlinear ultrasound

contrast becomes activated by protease activity. As representative targets, we selected the

constitutively active tobacco etch virus (TEV) endopeptidase, the calcium-dependent

mammalian protease calpain, and the processive bacterial protease ClpXP. We set out to test

the ability of acoustic biosensors engineered to respond to each of these enzymes to reveal

their activity under ultrasound, and to demonstrate biosensor imaging

in vitro

, in living

engineered cells, and

in vivo

in the mouse gastrointestinal (GI) tract.

RESULTS

Engineering an acoustic sensor of TEV endopeptidase

We selected the TEV endopeptidase as our first sensing target because of its well-

characterized recognition sequence and widespread use in biochemistry and synthetic

biology

. To sense TEV activity, we engineered a GvpC variant containing the TEV

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recognition motif ENLYFQ’G (Fig. 1b), hypothesizing that the cleavage of GvpC into two

smaller segments would cause the GV shell to become less stiff, thereby allowing it to

undergo buckling and produce enhanced nonlinear ultrasound contrast. We implemented this

design

in vitro

using GVs from

Anabaena flos-aque

(Ana), whose native GvpC can be

removed after GV isolation, and replaced with new versions expressed heterologously in

Escherichia coli

. Ana GvpC comprises five repeats of a predicted alpha-helical

polypeptide (Fig. 1a), and we tested insertions of the TEV recognition sequence, with and

without flexible linkers of different lengths, at several locations within this protein. After

incubating the engineered GVs with active TEV protease or a heat-inactivated “dead”

control (dTEV), we measured their hydrostatic collapse using pressurized absorbance

spectroscopy. This technique measures the optical density of GVs (which scatter 500 nm

light when intact) under increasing hydrostatic pressure, providing a quick assessment of

GV shell mechanics: GVs that collapse at lower pressures also produce more nonlinear

contrast

. Using this approach, we identified an engineered GV variant that showed

70 kPa reduction in its collapse pressure midpoint upon incubation with the active TEV

protease (Fig. 1c and Extended Data Fig. 1), and selected it for further characterization. This

GV sensor for TEV, hereafter referred to as GVS

TEV

, has the TEV cleavage site on the

second repeat of GvpC, flanked by flexible GSGSGSG linkers on both sides.

TEV cleavage of the GvpC on GVS

TEV

is expected to produce N- and C-terminal fragments

with molecular weights of approximately 9 and 14 kDa, respectively. Indeed, gel

electrophoresis of GVS

TEV

after exposure to active TEV resulted in the appearance of the

two cleaved GvpC fragments and a significant reduction in the intact GvpC band intensity

(Fig. 1d). In addition, removal from solution of unbound fragments via buoyancy

purification of the GVs resulted in a reduced band intensity for the N-terminal cleavage

fragment, indicating its partial dissociation after cleavage (Fig. 1d). No significant changes

in the GvpC band intensity were observed after incubation with dTEV. Transmission

electron microscopy (TEM) images showed intact GVs with similar appearance under both

conditions, confirming that protease cleavage did not affect the structure of the underlying

GV shell (Fig. 1e). Dynamic light scattering (DLS) showed no significant difference in the

hydrodynamic diameter of the engineered GVs after incubation with dTEV and active TEV

protease, confirming that the GVs remain dispersed in solution (Fig. 1f).

After confirming the desired mechanical and biochemical properties of GVS

TEV

, we imaged

it by ultrasound. Nonlinear imaging was performed in hydrogel samples containing the

biosensor, using a recently developed cross-amplitude modulation (x-AM) pulse sequence

x-AM uses pairs of cross-propagating plane waves to elicit highly specific nonlinear

scattering from buckling GVs at the wave intersection, while subtracting the linear signal

generated by transmitting each wave on its own

. Linear images were acquired using a

conventional B-mode sequence. As hypothesized, exposing the GVS

TEV

samples to TEV

protease produced a strong nonlinear acoustic response, with a maximal contrast-to-noise

ratio (CNR) enhancement of

7 dB at an applied acoustic pressure of 438 kPa (Fig. 1g).

Substantially less nonlinear contrast was observed in controls exposed to dTEV, while, as

expected, both samples produced similar linear scattering. Consistent with the pressure-

dependent mechanics of the GV shell, the differential nonlinear acoustic response of

GVS

TEV

became evident at pressures above 295 kPa, and kept increasing until 556 kPa, at

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which point the GVs began to collapse (Fig. 1h and Extended Data Fig. 1). As an additional

control, we found that GVs with the wild-type GvpC sequence (GV

) showed no

difference in their hydrostatic collapse pressure or nonlinear acoustic contrast in response to

TEV protease (Extended Data Fig. 1), and no wild-type GvpC cleavage was seen upon gel

electrophoresis (Extended Data Fig. 1). These results established GVS

TEV

as an acoustic

biosensor of the TEV protease enzyme, and additionally provided an experimental template

to develop additional sensors.

Engineering an acoustic sensor of calpain

After validating our basic acoustic biosensor design using the model TEV protease, we

examined its generalizability to other endopeptidases. As our second target, we selected the

calcium-dependent cysteine protease calpain, a mammalian enzyme with critical roles in a

wide range of cell types

–

. The two most abundant isoforms of this protease, known as μ-

calpain and m-calpain, are expressed in many tissues and involved in processes ranging from

neuronal synaptic plasticity to cellular senescence

. We designed an acoustic biosensor

of μ-calpain by inserting the

-spectrin-derived recognition sequence QQEVY’GMMPRD

into Ana GvpC (Fig. 2a). We screened several versions of GvpC incorporating this cleavage

sequence, flanked by GSG or GSGSG linkers, at different positions within the second helical

repeat. Pressurized absorbance spectroscopy performed in buffers with and without calpain

and Ca

allowed us to identify a GV sensor for calpain (GVS

calp

), showing an

approximately 50 kPa decrease in hydrostatic collapse pressure in the presence of the

enzyme and its ionic activator (Fig. 2b and Extended Data Fig. 2). Electrophoretic analysis

confirmed cleavage and partial dissociation of the cleaved fragments from the GV surface

(Extended Data Fig. 2), while TEM showed no change in GV morphology (Extended Data

Fig. 2).

Ultrasound imaging of GVS

calp

revealed a robust nonlinear acoustic response when both

calpain and calcium were present (Fig. 2, c, e, g), but not in negative controls lacking either

or both of these analytes. A slight clustering tendency of GVS

calp

nanostructures, which was

attenuated by incubation with activated calpain (Extended Data Fig. 2), resulted in a slightly

higher B-mode signal for the negative controls. However, this did not significantly affect the

maximal nonlinear sensor contrast of GVS

calp

of approximately 7dB (Fig. 2, c, e, g). This

contrast increased steeply beyond an applied acoustic pressure of 320 kPa (Fig. 2, d, f, h and

Extended Data Fig. 2). Using this biosensor, ultrasound imaging could be used to visualize

the dynamic response of calpain to Ca

, with a half-maximal response concentration of 140

μM (Fig. 2i and Extended Data Fig. 2). Additional control experiments performed on GVs

with wild-type GvpC showed no proteolytic cleavage, change in GV collapse pressure or

ultrasound response, after incubation with calcium-activated calpain (Extended Data Fig. 3).

These results show that acoustic biosensor designs based on GvpC cleavage can be

generalized to a mammalian protease and used to sense the dynamics of a conditionally

active enzyme.

Building an acoustic sensor of the protease ClpXP

In addition to endopeptidases, another important class of enzymes involved in cellular

protein signaling and homeostasis is processive proteases, which unfold and degrade full

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proteins starting from their termini

. To determine whether GV-based biosensors could be

developed for this class of enzymes, we selected ClpXP, a processive proteolytic complex

from

E. Coli

comprising the unfoldase ClpX and the peptidase ClpP

. ClpX recognizes and

unfolds protein substrates containing specific terminal peptide sequences called degrons.

The unfolded proteins are then fed into ClpP, which degrades them into small peptide

fragments

. We hypothesized that the addition of a degron to the C-terminus of GvpC

would enable ClpXP to recognize and degrade this protein, while leaving the underlying

GvpA shell intact, resulting in GVs with greater mechanical flexibility and nonlinear

ultrasound contrast (Fig. 3a).

To test this hypothesis, we appended the ssrA degron, AANDENYALAA, via a short SG

linker, to the C-terminus of Ana GvpC, resulting in a sensor that we named GVS

ClpXP

(Fig.

3a). We tested the performance of this biosensor

in vitro

using a reconstituted cell-free

transcription-translation system comprising

E. Coli

extract, purified ClpX, and a ClpP-

expressing plasmid. Gel electrophoresis performed after incubating GVS

ClpXP

with this cell-

free extract showed significant degradation of the engineered GvpC, compared to a negative

control condition in which the extract was pre-treated with a protease inhibitor (Fig. 3b).

TEM images showed intact GVs under both conditions, confirming that GvpC degradation

left the underlying GV shell uncompromised (Fig. 3c). Pressurized absorbance spectroscopy

indicated a substantial weakening of the GV shell upon ClpXP exposure, with the

hydrostatic collapse midpoint shifting by nearly 250 kPa (Fig. 3d and Extended Data Fig. 4).

Ultrasound imaging revealed a 17dB enhancement in the nonlinear contrast produced by

GVS

ClpXP

at an acoustic pressure of 477 kPa, in response to ClpXP activity (Fig. 3, e–f and

Extended Data Fig. 4). Control GVs containing wild type GvpC showed no sensitivity to

ClpXP (Fig. 3, g–i and Extended Data Fig. 4). These results establish the ability of GV-

based acoustic biosensors to visualize the activity of a processive protease as turn-on

sensors.

Constructing intracellular acoustic sensor genes

After demonstrating the performance of acoustic biosensors

in vitro

, we endeavored to show

that they could respond to enzymatic activity inside living cells. As the cellular host, we

chose

E. Coli

Nissle 1917. This probiotic strain of

E. Coli

has the capacity to colonize the

mammalian gastrointestinal tract, and is widely used as a chassis for the development of

microbial therapeutics

–

, making it a valuable platform for intracellular biosensors.

Recently, an engineered operon comprising GV-encoding genes from

Anabaena flos-aquae

and

Bacillus megaterium

was expressed in Nissle cells as acoustic reporter genes (

ARGs

allowing gene expression to be imaged with linear B-mode ultrasound

. To develop an

intracellular acoustic sensor gene targeting ClpXP (

ASG

ClpXP

), we swapped the wild type

gvpC

in the

ARG

gene cluster (

ARG

) with the modified

gvpC

from GVS

ClpXP

(dGvpC)

(Fig. 4a). For a first test of this intracellular biosensor, we transformed it into wild-type

(WT) Nissle cells, which natively express ClpXP protease, hypothesizing that it would show

a reduced intracellular collapse pressure and enhanced nonlinear contrast compared to

ARG

. Indeed, pressurized absorbance spectroscopy on intact cells expressing

ASG

ClpXP

revealed a reduction in the hydrostatic collapse pressure midpoint of

160 kPa relative to

cells expressing

ARG

(Extended Data Fig. 5). In ultrasound imaging, live cells

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expressing

ASG

ClpXP

showed an enhancement in nonlinear contrast of approximately 13 dB

(Extended Data Fig. 5), while linear B-mode signal was similar. The nonlinear response of

ASG

ClpXP

expressing cells was strongest beyond an acoustic pressure of 784 kPa (Extended

Data Fig. 5).

Next, to examine the ability of

ASG

ClpXP

to respond to intracellular enzymatic activity in a

dynamic manner, we generated a ClpXP-deficient strain of Nissle cells (Δ

clpXP

) through

genomic knock-out of the genes encoding ClpX and ClpP, and created a plasmid containing

these two genes under the control of an arabinose-inducible promoter (Fig. 4a). This allowed

us to externally control the activity of the ClpXP enzyme. Δ

clpXP

Nissle cells were co-

transformed with an inducible

clpX-clpP

(

clpXP

) plasmid and

ASG

ClpXP

. ClpXP production

in these cells after induction with L-arabinose resulted in an approximately 160 kPa

reduction in the hydrostatic collapse pressure midpoint (Fig. 4b and Extended Data Fig. 5).

Under ultrasound imaging, cells with induced ClpXP activity showed substantially stronger

nonlinear contrast (+6.7 dB) compared to cells uninduced for this protease (Fig. 4c), while

showing a similar B-mode signal. This enhancement in nonlinear signal was detectable with

acoustic pressures above 950 kPa (Fig. 4d and Extended Data Fig. 5). These experiments

demonstrate the ability of

ASG

ClpXP

to function as an intracellular acoustic sensor to

monitor variable enzyme activity.

A major application of dynamic sensors in cells is to monitor the activity of natural or

synthetic gene circuits

–

. To test if our acoustic sensors could be used to track the output

of a synthetic gene circuit in cells, we co-transformed WT Nissle cells with

ASG

ClpXP

, and a

separate wild-type

gvpC

gene controlled by anhydrotetracycline (aTc) (Fig. 4e). Our

hypothesis was that induction of this gene circuit only with IPTG would result in the

production of GVs with ClpXP-degradable GvpC, resulting in nonlinear contrast, whereas

the additional input of aTc would result in the co-production of non-degradable wild-type

GvpC, which would take the place of any degraded engineered GvpC on the biosensor shell

and lead to reduced nonlinear scattering (Fig. 4e). Indeed, when we induced cells with just

IPTG we observed strong nonlinear contrast. However, when aTc was added to the cultures

after IPTG induction, this contrast was reduced by approximately 10 dB (Fig. 4f–g and

Extended Data Fig. 5). These results, together with our findings in Δ

clpXP

cells with

inducible ClpXP, show that acoustic biosensors can be used to visualize the output of

synthetic gene circuits.

Ultrasound imaging of intracellular ClpXP activity in vivo

Finally, after establishing the basic principles of acoustic biosensor engineering

in vitro

and

demonstrating their performance in living cells, we assessed the ability of our sensor

constructs to produce ultrasound contrast within a biologically relevant anatomical location

in vivo

. In particular, approaches to imaging microbes in the mammalian GI tract

–

are

needed to support the study of their increasingly appreciated roles in health and disease

and the development of engineered probiotic agents

. The GI tract is also an excellent

target for ultrasound imaging due to its relatively deep location inside the animal, and the

use of ultrasound in clinical diagnosis and animal models of GI pathology, with appropriate

measures taken to minimize potential interference from air bubbles and solid matter

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To demonstrate the ability of acoustic biosensors to produce nonlinear ultrasound contrast

within the

in vivo

context of the mouse GI tract, we first co-injected WT Nissle cells

expressing

ASG

ClpXP

and

ARG

into the mouse colon (schematic shown in Extended

Data Fig. 6), distributing one cell population along the lumen wall and the other in the lumen

center. In these proof-of-concept experiments, the cells are introduced into the colon in a

rectally-injected agarose hydrogel to enable precise positioning and control over

composition. Using nonlinear ultrasound imaging, we could clearly visualize the unique

contrast generated by the protease-sensitive

ASGs

as a bright ring of contrast lining the

colon periphery (Fig. 5a). When the spatial arrangement was reversed, the bright nonlinear

contrast was concentrated in the middle of the lumen (Extended Data Fig. 7). A comparison

of ultrasound images acquired before and after acoustic collapse of the GVs, using a high-

pressure pulse from the transducer, confirmed that the bright ring of nonlinear contrast was

emanating from

ASG

ClpXP

-expressing cells (Fig. 5a), and this result was consistent across

independent experiments in 9 mice (Fig. 5b).

To demonstrate

in vivo

imaging of enzyme activity, we introduced Δ

clpXP

Nissle cells

expressing

ASG

ClpXP

into the mouse colon, with and without transcriptionally activating

intracellular ClpXP (schematic shown in Extended Data Fig. 6) . As above, the cells were

contained in an agarose hydrogel. Cells induced to express this enzyme showed enhanced

nonlinear contrast compared to cells not expressing ClpXP (Fig. 5c). Acoustic collapse

confirmed the acoustic biosensors as the primary source of nonlinear signal (Fig. 5c). This

performance was consistent across 7 mice and 2 spatial arrangements of the cells (Fig. 5d).

These results demonstrate the ability of acoustic biosensors to visualize enzyme activity

within the context of

in vivo

imaging.

Besides molecular sensing, one additional benefit of the nonlinear contrast generated by

ASG

ClpXP

-expressing cells is to make the cells easier to detect relative to background tissue

compared to linear B-mode imaging. Indeed, the nonlinear contrast of WT Nissle cells

expressing

ASG

ClpXP

had a significantly higher contrast-to-tissue ratio than either the

nonlinear contrast of

ARG

-expressing cells, or the B-mode contrast of either of these two

species (Extended Data Fig. 8).

DISCUSSION

Our results establish a paradigm for visualizing protease activity non-invasively with

ultrasound imaging. This paradigm is enabled by the dependence of the buckling mechanics

of GVs on the reinforcing protein GvpC, and the ability to turn this protein into a protease

substrate by incorporating specific internal or terminal peptide sequences. Similar to the

earliest work on fluorescent biosensors

, this initial study has focused on proteases due

to the importance of this class of enzymes in biology, their relatively compact recognition

motifs, and the large impact of their activity on protein structure. Based on our success in

sensing the function of three distinct proteases, we anticipate that the basic design strategy

presented here should be applicable to many enzymes of this type.

Our study lends itself to numerous future investigations to extend the applications of

acoustic protease sensors beyond the proof-of-concept demonstrations shown here. While

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our experiments in

E. Coli

and within the mouse GI tract establish the critical ability of such

biosensors to produce ultrasound contrast in relevant biological settings, additional

application-centric optimizations would enable the use of these constructs to address specific

problems in basic and synthetic biology. For example, purified acoustic biosensors could be

designed to sense extracellular proteases, which play homeostatic and disease-causing roles

in tissues ranging from extracellular matrix remodeling and blood clot formation to inter-

cellular signaling. Meanwhile, the expression of acoustic biosensor genes in cells could be

used to monitor natural cellular enzyme activity or serve as the output of synthetic signaling

pathways. Intracellular use in bacteria could be particularly relevant in studying microbes in

the mammalian GI tract, provided the successful adaptation of acoustic sensor genes to the

relevant host species and ensuring successful delivery via oral gavage, colonization and

metabolic viability. For potential applications in mammalian cells, acoustic protease sensor

designs must be integrated into recently developed genetic programs enabling the expression

of GVs in mammalian cells

. Successful use of acoustic sensors in this context will require

increasing the level of mammalian GV expression to enable non-destructive nonlinear

imaging.

In parallel, significant scope exists for further optimizing and generalizing the design of

acoustic biosensors. While all three of our sensors produced detectable nonlinear contrast in

response to protease activity, the changes exhibited by GVS

ClpXP

were significantly larger

than for the other two constructs. This is not surprising for an enzyme that processively

degrades GvpC, and whose recognition motif can be incorporated outside the main GV-

binding region of GvpC. Endopeptidase sensors could be optimized to reach similar

performance by incorporating more than one cleavage site within the GvpC sequence and

tuning the linkers connecting these sites to the rest of the protein. As with other protease

biosensors, the irreversibility of proteolysis means that for repeated or continuous sensing, it

is necessary for new sensor molecules to be synthesized or delivered. For genetically

encoded biosensors, this occurs through gene expression, potentially posing a metabolic

burden to the cell. For GVs, this burden could be reduced by re-expressing only the

engineered GvpC rather than the full GV, since this protein can be added onto the shell of

existing GVs, as demonstrated in this study and previous work

. Going beyond proteolytic

sensors, we anticipate that our biosensor design strategy could be modified to enable

allosteric conformational changes in GvpC, rather than its cleavage, to alter ultrasound

contrast, thereby creating acoustic biosensors that respond reversibly to non-cleaving

enzymes, ions or other signals of interest.

In addition to optimizing the biosensor constructs, it is also possible to improve the

ultrasound techniques used for their visualization. In this study, we monitored the activation

of our biosensors using a nonlinear x-AM pulse sequence, quantifying the resulting contrast

relative to linear B-mode scattering. This ratiometric signal is advantageous for

quantification in scenarios where the sensor concentration may vary. However, the

dependence of the x-AM response on applied acoustic pressure introduces a variable that

may differ across the ultrasonic field of view, and strategies involving dynamic pressure

adjustment may be needed to obtain the optimal signal from each point in the imaged plane.

In addition, normalization to B-mode signal in complex

in vivo

contexts may require

methods to separate the linear scattering contributions of acoustic sensors from those of

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background tissue. With these improvements, acoustic biosensors promise to take dynamic

imaging of molecular and cellular function to new depths.

ONLINE METHODS

Design and cloning of genetic constructs

All gene sequences were codon optimized for

E.Coli

expression and inserted into their

plasmid backbones via Gibson Assembly or KLD Mutagenesis using enzymes from New

England Biolabs and custom primers from Integrated DNA Technologies. The protease

recognition sequences for TEV protease and μ-calpain, flanked by flexible linkers, were

introduced by substitution-insertion into the second repeat of the wild-type Ana

gvpC

gene

sequence in a pET28a expression vector (Novagen) driven by a T7 promoter and lac

operator. The ssrA degradation tag for the ClpXP bacterial proteasome was appended to the

C-terminus of Ana

gvpC

using a short flexible linker. The acoustic sensor gene for

intracellular protease sensing of ClpXP was constructed by modifying of the acoustic

reporter gene cluster

ARG1

, by addition of the ssrA degradation tag to the C-terminal of

gvpC

using a linker sequence. For expression in

E.Coli

Nissle 1917 cells, the pET28a T7

promoter was replaced by the T5 promoter. For inducible expression of

clpX

and

clpP

, the

genes encoding those two proteins were cloned from the

E. Coli

Nissle 1917 genome into a

modified pTARA backbone under a P

BAD

promoter and araBAD operon. For dynamic

regulation of intracellular sensing, the wild-type GvpC sequence was cloned into a modified

pTARA backbone under a pTet promoter and tetracycline operator. The complete list and

source of plasmids used in this study is given in Supplementary Table 1. Plasmid constructs

were cloned using NEB Turbo

E. Coli

(New England Biolabs) and sequence-validated.

Construction of

clpX

−

clpP

−

strain of

E.Coli

Nissle 1917 (Δ

clpXP

)

The knockout of

clpX

and

clpP

E.Coli

Nissle (ECN) was accomplished by Lambda Red

recombineering using previously published methods

. A FRT-flanked

cat

gene was

recombined into ECN genome to replace the

clpX

and

clpP

genes, and the integrated cat

gene was then removed by the

FLP

recombinase from pE-FLP

to yield the Δ

clpXP

strain.

More information on the recombineering plasmids used in this study and their source is

provided in Supplementary Table 1.

GV expression, purification and quantification

For

in vitro

assays, GVs were harvested and purified from confluent

Ana

cultures using

previously published protocols

. Briefly, Ana cells were grown in Gorham’s media

supplemented with BG-11 solution (Sigma) and 10 mM sodium bicarbonate at 25°C, 1%

and 100 rpm shaking, under a 14h light and 10h dark cycle. Confluent cultures were

transferred to sterile separating funnels and left undisturbed for 2–3 days to allow buoyant

Ana

cells expressing GVs to float to the top and for their subnatant to be drained.

Hypertonic lysis with 10% Solulyse (Genlantis) and 500 mM sorbitol was used to release

and harvest the Ana GVs. Purified GVs were obtained through 3–4 rounds of centrifugally

assisted floatation, with removal of the subnatant and resuspension in phosphate buffered

saline (PBS, Corning) after each round.

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For expression of acoustic reporter/sensor genes (

ARG/ASG

) in bacteria, wild-type

E. Coli

Nissle 1917 cells (Ardeypharm GmbH) were made electrocompetent and transformed with

the genetic constructs. After electroporation, cells were rescued in SOC media supplemented

with 2% glucose for 1h at 37°C. Transformed cells were grown for 12–16 hours at 37°C in 5

mL of LB medium supplemented with 50 μg/mL kanamycin and 2% glucose. Large-scale

cultures for expression were prepared by a 1:100 dilution of the starter culture in LB

medium containing 50 μg/mL kanamycin and 0.2% glucose. Cells were grown at 37°C to an

600nm

of 0.2–0.3, then induced with 3μM Isopropyl

-D-1-thiogalactopyranoside (IPTG)

and allowed to grow for 22 hrs at 30°C. Buoyant

E.Coli

Nissle cells expressing GVs were

isolated from the rest of the culture by centrifugally assisted floatation in 50 mL conical

tubes at 300g for 3–4 hrs, with a liquid column height less than 10 cm to prevent GV

collapse by hydrostatic pressure.

The concentration of Ana GVs was determined by measurement of their optical density

(OD) at 500 nm (OD

500

) using a Nanodrop spectrophotometer (Thermo Fisher Scientific),

using the resuspension buffer or collapsed GVs as the blank. As established in previous

work

, the concentration of GVs at OD

500

= 1 is approximately 114 pM and the gas

fraction is 0.0417%. The OD of buoyant cells expressing GVs were quantified at 600 nm

using the Nanodrop.

Bacterial expression and purification of GvpC variants

For expression of Ana GvpC variants, plasmids were transformed into chemically competent

BL21(DE3) cells (Invitrogen) and grown overnight for 14–16 h at 37°C in 5 mL starter

cultures in LB medium with 50 μg/mL kanamycin. Starter cultures were diluted 1:250 in

Terrific Broth (Sigma) and allowed to grow at 37°C (250 rpm shaking) to reach an OD

600nm

of 0.4–0.7. Protein expression was induced by addition of 1 mM IPTG, and the cultures

were transferred to 30°C. Cells were harvested by centrifugation at 5500g after 6–8 hours.

For the GvpC-ssrA variant, expression was carried out at 25°C for 8 hours to reduce the

effect of protease degradation and obtain sufficient protein yield.

GvpC was purified from inclusion bodies by lysing the cells at room temperature using

Solulyse (Genlantis), supplemented with lysozyme (400 μg/mL) and DNase I (10 μg/mL).

Inclusion body pellets were isolated by centrifugation at 27,000g for 15 mins and then

resuspended in a solubilization buffer comprising 20 mM Tris-HCl buffer with 500 mM

NaCl and 6 M urea (pH: 8.0), before incubation with Ni-NTA resin (Qiagen) for 2 h at 4°C.

The wash and elution buffers were of the same composition as the solubilization buffer, but

with 20mM and 250 mM imidazole respectively. The concentration of the purified protein

was assayed using the Bradford Reagent (Sigma). Purified GvpC variants were verified to be

>95% pure by SDS-PAGE analysis.

Preparation of gas vesicles for

in vitro

protease assays

Engineered GVs having protease-sensitive or wild-type GvpC were prepared using urea

stripping and GvpC re-addition

. Briefly, Ana GVs were stripped of their native outer

layer of GvpC by treatment with 6M urea solution buffered with 100 mM Tris- HCl (pH:8–

8.5). Two rounds of centrifugally assisted floatation with removal of the subnatant liquid

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after each round were performed to ensure complete removal of native GvpC. Recombinant

Ana GvpC variants purified from inclusion bodies were then added to the stripped Ana GVs

in 6 M urea a 2–3x molar excess concentration determined after accounting for 1:25 binding

ratio of GvpC: GvpA. For a twofold stoichiometric excess of GvpC relative to binding sites

on an average Ana GV, the quantity of recombinant GvpC (in nmol) to be added to stripped

GVs was calculated according to the formula: 2 * OD * 198 nM * volume of GVs (in liters).

The mixture of stripped GVs (OD

500nm

= 1–2) and recombinant GvpC in 6 M urea buffer

was loaded into dialysis pouches made of regenerated cellulose membrane with a 6–8 kDa

M.W. cutoff (Spectrum Labs). The GvpC was allowed to slowly refold onto the surface of

the stripped GVs by dialysis in 4 L PBS for at least 12 h at 4 °C. Dialyzed GV samples were

subjected to two or more rounds of centrifugally assisted floatation at 300 g for 3–4 h to

remove any excess unbound GvpC. Engineered GVs were resuspended in PBS after

subnatant removal and quantified using pressure-sensitive OD measurements at 500 nm

using a Nanodrop.

Pressurized absorbance spectroscopy

Purified, engineered Ana GVs were diluted in experimental buffers to an OD

500nm

0.2–0.4,

and 400 μL of the diluted sample was loaded into a flow-through quartz cuvette with a

pathlength of 1 cm (Hellma Analytics). Buoyant

E.Coli

Nissle cells expressing GVs were

diluted to an OD

600nm

1 in PBS for measurements. A 1.5 MPa nitrogen gas source was

used to apply hydrostatic pressure in the cuvette through a single valve pressure controller

(PC series, Alicat Scientific), while a microspectrometer (STS-VIS, Ocean Optics) measured

the OD of the sample at 500 nm (for Ana GVs) or 600 nm (for Nissle cells). The hydrostatic

pressure was increased from 0 to 1 MPa in 20 kPa increments with a 7 second equilibration

period at each pressure before OD measurement. Each set of measurements was normalized

by scaling to the Min-Max measurement value, and the data was fitted using the Boltzmann

sigmoid function

= 1 +

−

ΔP

−1

, with the midpoint of normalized OD change (P

)

and the 95% confidence intervals, rounded to the nearest integer, reported in the figures.

TEM sample preparation and imaging

Freshly diluted samples of engineered Ana GVs (OD

500nm

0.3) in 10 mM HEPES buffer

containing 150 mM NaCl (pH 8) were used for TEM. 2 μL of the sample was added to

Formvar/carbon 200 mesh grids (Ted Pella) that were rendered hydrophilic by glow

discharging (Emitek K100X). 2% uranyl acetate was added for negative staining. Images

were acquired using the FEI Tecnai T12 LaB6 120kV TEM equipped with a Gatan Ultrascan

2k X 2k CCD and ‘Leginon’ automated data collection software suite.

Dynamic light scattering (DLS) measurements

Engineered Ana GVs were diluted to an OD

500nm

0.2 in experimental buffers. 150–200 μL

of the sample was loaded into a disposable cuvette (Eppendorf UVette®) and the particle

size was measured using the ZetaPALS particle sizing software (Brookhaven instruments)

with an angle of 90 ° and refractive index of 1.33.

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Denaturing polyacrylamide gel electrophoresis (SDS-PAGE)

GV samples were OD

500nm

matched and mixed 1:1 with 2x Laemmli buffer (Bio-Rad),

containing SDS and 2-mercaptoethanol. The samples were then boiled at 95°C for 5 minutes

and loaded into a pre-made polyacrylamide gel (Bio-Rad) immersed in 1x Tris-Glycine-SDS

Buffer. 10 uL of Precision Plus Protein™ Dual Color Standards (Bio-Rad) was loaded as the

ladder. Electrophoresis was performed at 120V for 55 minutes, after which the gel was

washed in DI water for 15 minutes to remove excess SDS and commassie-stained for 1 hour

in a rocker-shaker using the SimplyBlue SafeStain (Invitrogen). The gel was allowed to

destain overnight in DI water before imaging using a Bio-Rad ChemiDoc™ imaging system.

In vitro

protease assays

For

in vitro

assays with the TEV endopeptidase, recombinant TEV protease (R&D Systems,

Cat. No. 4469-TP-200) was incubated (25% v/v fraction) with engineered Ana GVs

resuspended in PBS (final OD

500nm

in reaction mixture = 5–6) at 30°C for 14–16 h. This

corresponds to a TEV concentration of 0.1

0.125 mg/mL (depending on the lot), within the

range used in previous studies with this enzyme

. Engineered GVs with wild-type GvpC

and TEV protease heat-inactivated at 80°C for 20–30 mins were used as the controls.

For

in vitro

assays with calpain, calpain-1 from porcine erythrocytes (Millipore Sigma, Cat.

No. 208712) was incubated in a 10% v/v fraction with engineered Ana GVs in a reaction

mixture containing 50 mM Tris-HCl, 50 mM NaCl, 5 mM 2-mercaptoethanol, 1 mM EDTA

and 1 mM EGTA and 5 mM Ca

(pH: 7.5) This corresponds to a calpain concentration of ≥

0.168 units per μl, with 1 unit defined by the manufacturer as sufficient to cleave 1 pmol of a

control fluorogenic substrate in 1 min at 25°C. The final concentration of engineered GVs in

the reaction mixture was OD

500nm

6 and the protease assay was carried out at 25°C for 14–

16h. Negative controls included the same reaction mixture without calpain, without calcium,

or without calpain and calcium. Engineered GVs with WT-GvpC were used as additional

negative controls.

For

in vitro

assays with ClpXP, a reconstituted cell-free transcription-translation (TX-TL)

system adapted for ClpXP degradation assays

(gift from Zachary Sun and Richard

Murray) was used. Briefly, cell-free extract was prepared by lysis of ExpressIQ

E.Coli

cells

(New England Biolabs), and mixed in a 44% v/v ratio with an energy source buffer, resulting

in a master mix of extract and buffer comprising: 9.9 mg/mL protein, 1.5 mM each amino

acid except leucine, 1.25 mM leucine, 9.5 mM Mg-glutamate, 95 mM K-glutamate, 0.33

mM DTT, 50 mM HEPES, 1.5 mM ATP and GTP, 0.9 mM CTP and UTP, 0.2 mg/mL tRNA,

0.26 mM CoA, 0.33 mM NAD, 0.75 mM cAMP, 0.068 mM folinic acid, 1 mM spermidine,

30 mM 3-PGA and 2% PEG-8000. For purified ClpX protein, a monomeric N-terminal

deletion variant Flag-ClpXdeltaNLinkedHexamer-His6

(Addgene ID: 22143) was used.

Post Ni-NTA purification, active fractions of ClpX hexamers with sizes above 250 kDa were

isolated using a Supradex 2010/300 column, flash frozen at a concentration of 1.95 μM and

stored at −80°C in a storage buffer consisting of: 50 mM Tris-Cl (pH 7.5), 100 mM NaCl,

1mM DTT, 1 mM EDTA and 2% DMSO. The final reaction mixture was prepared as

follows: 75% v/v fraction of the master mix, 10% v/v of purified ClpX, 1nm of the purified

pBEST-ClpP plasmid and engineered Ana GVs (concentration of OD

500nm

= 2.5–2.7 in the

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reaction mixture). The mixture was made up to the final volume using ultrapure H

O. The

reaction was allowed to proceed at 30°C for 14–16 h. As a negative control, a protease

inhibitor cocktail mixture (SIGMA

FAST

™, Millipore Sigma) was added to the reaction

mixture at 1.65x the manufacturer-recommended concentration and pre-incubated at room

temperature for 30 mins.

Dynamic sensing of ClpXP activity in Δ

clpXP E.Coli

Nissle 1917 cells

ClpXP

E. Coli

Nissle 1917 cells were made electrocompetent and co-transformed with the

pET expression plasmid (Lac-driven) containing the

ASG

for ClpXP and a modified pTARA

plasmid (pBAD-driven) containing the

clpX

and

clpP

genes. Electroporated cells were

rescued in SOC media supplemented with 2% glucose for 2h at 37°C. Transformed cells

were grown overnight at 37°C in 5 mL LB medium supplemented with 50 μg/mL

kanamycin, 25 μg/mL chloramphenicol and 2% glucose. Starter cultures were diluted 1:100

in LB medium with 50 μg/mL kanamycin, 25 μg/mL chloramphenicol and 0.2% glucose and

allowed to grow at 37 °C to reach an OD

600nm

of 0.2–0.3.

ASG

expression was induced with

3μM IPTG and the bacterial culture was transferred to the 30 °C incubator with 250 rpm

shaking for 30 minutes. The culture was then split into two halves of equal volume, and one

half was induced with 0.5% (weight fraction) L-arabinose for expression of ClpXP protease.

Cultures with and without L-arabinose induction were allowed to grow for an additional 22

h at 30°C. Cultures were then spun down at 300 g in a refrigerated centrifuge at 4 °C for 3–4

h in 50 mL conical tubes to isolate buoyant cells expressing GVs from the rest of the culture.

The liquid column height was maintained at less than 10 cm to prevent GV collapse by

hydrostatic pressure.

Dynamic sensing of circuit-driven gene expression in

E.Coli

Nissle 1917 cells

Electrocompetent

E. Coli

Nissle cells were co-transformed with the pET expression plasmid

(Lac-driven) containing the ASG for ClpXP and a modified pTARA plasmid

(Tet-driven)

containing the WT Ana GvpC gene. Electroporated cells were rescued in SOC media

supplemented with 2% glucose for 2h at 37°C. Transformed cells were grown overnight at

37°C in 5 mL LB medium supplemented with 50 μg/mL kanamycin, 50 μg/mL

chloramphenicol and 2% glucose. Starter cultures were diluted 1:100 in LB medium with 50

μg/mL kanamycin, 50 μg/mL chloramphenicol and 0.2% glucose and allowed to grow at 37

°C to reach an OD

600nm

of 0.2–0.3. ASG expression was induced with 3 μM IPTG and the

bacterial culture was transferred to 30 °C incubator with 250 rpm shaking for 1.5–2 h. The

culture was then split into two halves of equal volume, and one half was induced with 50

ng/mL aTc for expression of WT GvpC. Cultures with and without aTc induction were

allowed to grow for an additional 20 h at 30°C. Cultures were then spun down at 300 g in a

refrigerated centrifuge at 4 °C for 3–4 h in 50 mL conical tubes to isolate buoyant cells

expressing GVs from the rest of the culture. The liquid column height was maintained at less

than 10 cm to prevent GV collapse by hydrostatic pressure.

In vitro

ultrasound imaging

Imaging phantoms were prepared by melting 1% agarose (w/v) in PBS and casting wells

using a custom 3-Dprinted template mold containing a 2-by-2 grid of cylindrical wells with

2 mm diameter and 1 mm spacing between the outer radii in the bulk material. Ana GV

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samples from

in vitro

assays or buoyant Nissle cells expressing GVs were mixed 1:1 with

1% molten agarose solution at 42°C and quickly loaded before solidification into the

phantom wells. All samples and their controls were OD-matched using the Nanodrop prior

to phantom loading, with the final concentration being OD

500nm

= 2.2 for Ana GVs and

600nm

= 1.0–1.5 for buoyant Nissle cells. Wells not containing sample were filled with

plain 1% agarose. Hydrostatic collapse at 1.4 MPa was used to determine that the

contribution to light scattering from GVs inside the cells was similar for those expressing the

acoustic sensor gene and its wild-type

ARG

counterpart. The phantom was placed in a

custom holder on top of an acoustic absorber material and immersed in PBS to acoustically

couple the phantom to the ultrasound imaging transducer.

Imaging was performed using a Verasonics Vantage programmable ultrasound scanning

system and a L22–14v 128-element linear array Verasonics transducer, with a specified pitch

of 0.1 mm, an elevation focus of 8 mm, an elevation aperture of 1.5mm and a center

frequency of 18.5 MHz with 67% −6 dB bandwidth. Linear imaging was performed using a

conventional B-mode sequence with a 128-ray-lines protocol. For each ray line, a single

pulse was transmitted with an aperture of 40 elements. For nonlinear image acquisition, a

custom cross-amplitude modulation (x-AM) sequence detailed in an earlier study

, with an

x-AM angle (

) of 19.5° and an aperture of 65 elements, was used. Both B-mode and x-AM

sequences were programmed to operate close to the center frequency of the transducer

(15.625 MHz) and the center of the sample wells were aligned to the set transmit focus of 5

mm. Transmitted pressure at the focus was calibrated using a Precision Acoustics fiber-optic

hydrophone system. Each image was an average of 50 accumulations. B-mode images were

acquired at a transmit voltage of 1.6V (132 kPa), and an automated voltage ramp imaging

script (programmed in MATLAB) was used to sequentially toggle between B-mode and x-

AM acquisitions. The script acquired x-AM signals at each specified voltage step,

immediately followed by a B-mode acquisition at 1.6V (132 kPa), before another x-AM

acquisition at the next voltage step. For engineered Ana GVs subjected to

in vitro

protease

assays, an x-AM voltage ramp sequence from 4V (230 kPa) to 10V (621 kPa) in 0.2V

increments was used. For wild-type Nissle cells expressing GVs, an x-AM voltage ramp

sequence from 7.5V (458 kPa) to 25V (1.6 MPa) in 0.5V increments was used. Samples

were subjected to complete collapse at 25V with the B-mode sequence for 10 seconds, and

the subsequent B-mode image acquired at 1.6V and x-AM image acquired at the highest

voltage of the voltage ramp sequence was used as the blank for data processing. There was

no significant difference between the signals acquired at specific acoustic pressures during a

voltage ramp or after directly stepping to the same pressure (Extended Data Fig. 9).

Due to transducer failure, a replacement Verasonics transducer (L22–14vX) with similar

specifications was used in experiments with Δ

clpXP

cells. The transmitted pressure at the

focus was calibrated in the same way as the L22–14v. B-mode images were acquired at a

transmit voltage of 1.6V (309 kPa), and an x-AM voltage ramp sequence from 6V (502 kPa)

to 25V (2.52 MPa) was used. The imaging protocol was otherwise unchanged.

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In vivo

ultrasound imaging

All

in vivo

experiments were performed on C57BL/6J male mice, aged 14–34 weeks, under

a protocol approved by the Institutional Animal Care and Use Committee of the California

Institute of Technology. No randomization or blinding were necessary in this study. Mice

were anesthetized with 1–2% isoflurane, maintained at 37 °C on a heating pad, depilated

over the imaged region, and enema was performed by injecting PBS to expel gas and solid

contents in mice colon. For imaging of

E. Coli

in the gastrointestinal tract, mice were placed

in a supine position, with the ultrasound transducer positioned on the lower abdomen,

transverse to the colon such that the transmit focus of 5 mm was close to the center of the

colon lumen. Prior to imaging, two variants of buoyancy-enriched

E. Coli

Nissle 1917 were

mixed in a 1:1 ratio with 4% agarose in PBS at 42 °C, for a final bacterial concentration of

1.5E9 cells ml

−1

. An 8-gauge gavage needle was filled with the mixture of agarose and

bacteria of one cell population. Before it solidified, a 14-gauge needle was placed inside the

8-gauge needle to form a hollow lumen within the gel. After the agarose-bacteria mixture

solidified at room temperature for 10 min, the 14-gauge needle was removed. The hollow

lumen was then filled with the agarose–bacteria of the other cell population. After it

solidified, the complete cylindrical agarose gel was injected into the colon of the mouse with

a PBS back-filled syringe. For the colon imaging, imaging planes were selected to avoid gas

bubbles in the field of view. In all

in vivo

experiments, three transducers were used,

including two L22–14v and one L22–14vX, due to transducer failures unrelated to this

study. B-mode images were acquired at 1.9V (corresponding to 162 kPa in water) for L22–

14v, and 1.6V (309 kPa in water) for L22–14vX. x-AM images were acquired at 20V (1.27

MPa in water) for L22–14v and 15V (1.56 MPa in water) for L22–14vX, with other

parameters being the same as those used for

in vitro

imaging. B-mode anatomical imaging

was performed at 7.4V using the ‘L22–14v WideBeamSC’ script provided by Verasonics.

Image processing and data analysis

All

in vitro

and

in vivo

ultrasound images were processed using MATLAB. Regions of

interest (ROIs) were manually defined so as to adequately capture the signals from each

sample well or region of the colon. The sample ROI dimensions (1.2 mm × 1.2 mm square)

were the same for all

in vitro

phantom experiments. The noise ROI was manually selected

from the background for each pair of sample wells. For the

in vivo

experiments, circular

ROIs were manually defined to avoid edge effects from the skin or colon wall, and the tissue

ROIs were defined as the rest of the region within the same depth range of the signal ROIs.

For each ROI, the mean pixel intensity was calculated, and the pressure-sensitive ultrasound

intensity (Δ

intact

−

collapsed

) was calculated by subtracting the mean pixel intensity of

the collapsed image from the mean pixel intensity of the intact image. The contrast-to-noise

ratio (CNR) was calculated for each sample well by taking the mean intensity of the sample

ROI over the mean intensity of the noise ROI. The x-AM by B-mode ratio at a specific

voltage (or applied acoustic pressure) was calculated with the following formula:

X‐AM

B‐mode

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where ΔI

x−AM

(V) is the pressure-sensitive nonlinear ultrasound intensity acquired by the x-

AM sequence at a certain voltage V, and ΔI

B−mode

(V) is the pressure-sensitive linear

ultrasound intensity of the B-mode acquisitions at 1.6V (132 kPa) following the x-AM

acquisitions at the voltage V. All images were pseudo-colored (bone colormap for B-mode

images, hot colormap for x-AM images), with the maximum and minimum levels indicated

in the accompanying color bars.

Statistical analysis

Data is plotted as the mean ± standard error of the mean (SEM). Sample size is N=3

biological replicates in all

in vitro

experiments unless otherwise stated. For each biological

replicate, there were technical replicates to accommodate for variability in experimental

procedures such as sample loading and pipetting. SEM was calculated by taking the values

for the biological replicates, each of which was the mean of its technical replicates. The

numbers of biological and technical replicates were chosen based on preliminary

experiments such that they would be sufficient to report significant differences in mean

values. Individual data for each replicate is given in Extended Data Figures 1–9 in the form

of scatter plots. P values, for determining the statistical significance for the

in vivo

data,

were calculated using a two-tailed paired t-test.

Data availability

The authors declare that data supporting the findings of this study are available within the

article and its Supplementary Information. Additional data are available from the

corresponding author upon reasonable request.

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Extended Data

Extended Data Fig. 1. Engineering an acoustic sensor of TEV endopeptidase activity.

(a)

Coomassie-stained SDS-PAGE gel of OD

500nm

-matched samples of GV

incubated

with dTEV and TEV protease, before and after buoyancy purification (labeled pre b.p. and

post b.p., respectively). N = 3 biological replicates.

(b)

Scatter plots showing normalized

500nm

of GVS

TEV

as a function of hydrostatic pressure. (N = 3 biological replicates for

GVS

TEV

+ TEV and N =4 for GVS

TEV

+ dTEV.)

(c)

Scatter plots showing the ratio of

nonlinear (x-AM) to linear (B-mode) ultrasound signal as a function of applied acoustic

pressure for all the replicate samples used in the x-AM voltage ramp imaging experiments

for GVS

TEV

. N = 3 biological replicates and total number of replicates is 8.

(d)

Scatter plots

showing normalized OD

500nm

of GV

as a function of hydrostatic pressure. (N = 3

biological replicates for GV

+dTEV and N=4 for GV

+ TEV.)

(e)

Representative

ultrasound images of agarose phantoms containing GV

incubated with TEV or dTEV

protease at OD

500nm

2.2. The B-mode image was acquired at 132kPa and the x-AM image at

569 kPa. Similar images acquired for N=3 biological replicates, with each N consisting of 3

technical replicates. CNR stands for contrast-to-noise-ratio, and color bars represent relative

ultrasound signal intensity on the dB scale. Scale bars represent 1 mm

(f)

Scatter plots

showing the ratio of nonlinear (x-AM) to linear (B-mode) ultrasound signal as a function of

applied acoustic pressure for all the replicate samples used in the x-AM voltage ramp

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imaging experiments for GV

. N=3 biological replicates, with each N consisting of 3

technical replicates. Solid curve represents the mean of all the replicates.

Extended Data Fig. 2. Engineering an acoustic sensor of calpain activity.

(a)

Individual scatter plots for Fig. 2(b). N = 5 biological replicates for +Calp/+Ca

, 6 for

−Calp/+Ca

and +Calp/−Ca

, 7 for −Calp/−Ca

(b)

Coomassie-stained SDS-PAGE gel

of OD

500nm

-matched samples of GVS

calp

incubated in the presence (+) or absence (−) of

calpain (first +/−) and calcium (second +/−), before and after buoyancy purification (labeled

pre b.p. and post b.p. respectively). N = 3 biological replicates.

(c)

Representative TEM

images of GVS

calp

after incubations in the presence or absence of calpain and/or calcium.

Scale bars represent 100 nm. At least 20 GV particles were imaged for each condition.

(d)

DLS measurements showing the average hydrodynamic diameter of GVS

calp

and GV

samples after calpain/calcium incubations (N = 2 biological replicates for GVS

calp

+/−, +/+,

+/+ and 3 for other conditions, individual dots represent each N and horizontal line

indicates the mean). Error bars indicate SEM when N = 3. (

e, f, g

) Individual scatter plots

for Fig. 2(d, f, h). N = 3 biological replicates with each N consisting of 2 technical replicates

(total number of replicates is 18 for +/+ and 6 for each of the remaining conditions). Solid

line represents the mean of all the replicates for (

a, e–g

). (h) Scatter plots for Fig. 2i; N = 3

biological replicates, individual dots represent each N and solid blue line showing the fitted

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curve (a Hill equation with a coefficient of 1, with a half-maximum response concentration

(EC

) of 140 μm).

Extended Data Fig. 3. Characterization of GV

sample with calpain protease.

(

a, b, c

) Representative ultrasound images of agarose phantoms containing GV

incubated

in the presence (+) or absence (−) of calpain (first +/−) and calcium (second +/−), at

500nm

2.2. The B-mode images were taken at 132 kPa for

a, b

and

and the x-AM

images corresponding to the maximum difference in non-linear contrast between the +/+

sample and the negative controls were taken at 438 kPa for

and

and at 425 kPa for

CNR stands for contrast-to-noise-ratio and color bars represent ultrasound signal intensity in

the dB scale. Scale bars represent 1 mm. N = 2 biological replicates for

a, b

and

. (

d, e, f

)

Scatter plots showing the ratio of x-AM to B-mode ultrasound signal as a function of

increasing acoustic pressure for GV

after incubation in the presence or absence of calpain

and/or calcium (N = 2 biological replicates). (

) Hydrostatic collapse curves of GV

after

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