Cell-selective proteomics for biological discovery
Shannon E. Stone
,
Weslee S. Glenn
,
Graham D. Hamblin
, and
David A. Tirrell
Division of Chemistry and Chemical Engineering, California Institute of Technology, Pasadena,
CA 91125
Abstract
Cells alter the proteome to respond to environmental and developmental cues. Global analysis of
proteomic responses is of limited value in heterogeneous environments, where there is no
“average” cell. Advances in sequencing, protein labeling, mass spectrometry, and data analysis
have fueled recent progress in the investigation of specific subpopulations of cells in complex
systems. Here we highlight recently developed chemical tools that enable cell-selective proteomic
analysis of complex biological systems, from bacterial pathogens to whole animals.
Graphical Abstract
Keywords
cell-selective; proteome; TRAP; CTAP; BONCAT; SORT; OP-Puro; APEX; newly-synthesized;
subcellular; click chemistry
Introduction
Cellular protein synthesis changes rapidly in response to internal and external cues in ways
that vary from cell to cell. Global proteomic analyses of microbial communities, tissues and
organisms have provided important insights into the behavior of such systems, but can
obscure the diversity of responses characteristic of different cellular subpopulations (Figure
1). Cell-selective methods for the analysis of protein synthesis are being developed to
resolve proteomic changes in space and time.
Cell-type-specific transcriptomics experiments have revealed mRNA expression patterns in a
wide array of biological systems, but mRNA and protein levels are often dissonant [
1
].
Corresponding Author:
David A. Tirrell, tirrell@caltech.edu.
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Moreover, some important elements of proteome dynamics, including posttranslational
modification, degradation, and localization, cannot be addressed by mRNA measurements
alone [
2
,
3
]. Until recently, changes in protein abundance in specific cells could be measured
only in targeted, low-throughput experiments, but innovations in mass spectrometry and
computational algorithms have facilitated the identification and quantification of thousands
of proteins simultaneously from complex biological samples [
4
–
6
].
In this Opinion, we highlight recent developments in determining cell-type-specific
proteomes and recommend experimental design strategies that are guided by the question at
hand.
Cell-selective translatomics and ribosome profiling
Translatomic studies, which select for ribosome-associated transcripts, have yielded stronger
correlations between transcript and protein abundances than experiments that measure
steady-state mRNA levels [
7
]. Cell-type-specific studies have been enabled by translating
ribosome affinity purification (TRAP), a method in which epitope-tagged ribosomes and
their associated transcripts are captured, enriched and subjected to amplification and deep
sequencing [
8
]. TRAP can be rendered cell-specific by placing expression of the tagged
ribosome under control of a selective promoter.
More recently, Ingolia and Weissman have developed ribosome profiling, which identifies
ribosome-protected mRNA footprints and allows investigators to determine ribosome
occupancy with positional specificity. This information can be used to measure translation
levels and locate non-canonical start sites [
7
]. Gonzalez
et al
. used TRAP to cell-selectively
purify ribosome-bound transcripts, and employed ribosome profiling to identify the
translatome of gliomas and to reveal decreased translation in glial progenitors compared to
the tumor microenvironment [
9
]. Ribosome profiling is a powerful technique that we expect
to find increasing use upon further development of cell-specific methods.
While translatomic studies provide greater depth of coverage than current proteomic
measurements, ribosome binding does not ensure that a transcript is undergoing active
translation [
10
].
Separating cells for steady-state proteomic analysis
The earliest strategies to determine cell-specific proteomes relied on separating and
purifying the cells of interest prior to analysis. Cells can be sorted on the basis of expression
of a transgene under control of a cell-specific promoter or by antibody staining of marker
epitopes. These tools are well established and have been thoughtfully reviewed [
10
,
11
].
Physical methods have been used for years to isolate cell types from mammalian tissues for
subsequent downstream analyses [
12
,
13
]. More recently these methods have been used to
measure growth rates and elucidate proteomic signatures of
Salmonella
during murine
infection [
14
].
Physical separations remain the best method for analyzing clinical specimens and
genetically intractable organisms. However, imperfect separations and long sample
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processing times can diminish selectivity and increase the likelihood of artifacts.
Furthermore, such methods intrinsically yield steady-state proteomic information. In
contrast, metabolic labeling strategies enable cell-specific proteomic analysis to be
accomplished in time-resolved fashion.
Metabolic labeling: trade-offs between sensitivity and perturbation
Metabolic labeling methods are temporally resolved and use an arsenal of amino acid
isotopologs, non-canonical amino acids, and analogs of protein synthesis inhibitors (Figure
2). Each of these strategies can be placed under control of cell-specific genetic elements to
afford cellular resolution. The choice of promoter(s) is key for these systems, and the degree
of protein labeling needs to be weighed against the possibility of perturbing the system.
Results should be validated via independent assays because labels may affect protein
expression, stability, and/or function.
Cell-type-specific labeling using amino acid precursors (CTAP)
Stable isotope labeling by amino acids in cell culture (SILAC) relies on the incorporation of
isotopically labeled amino acids into proteins. To make SILAC cell-selective, Gauthier
et al
.
introduced cell-type-specific labeling using amino acid precursors (CTAP), a method that
exploits the fact that lysine is an essential amino acid in mammalian cells [
15
]. Cell-selective
expression of biosynthetic enzymes allows L-lysine isotopologs to be synthesized
in situ
starting from isotope-labeled precursors. Only minor differences in gene expression resulted
from feeding the heavy precursor to cells expressing the biosynthetic machinery versus
supplementing cells directly with L-lysine.
In principle, both exchange of L-lysine between cells and extracellular processing of the
precursor can compromise the cell-specificity of the CTAP method. When Lavis and
coworkers employed an analogous strategy to unmask fluorophores in targeted cells, they
noted that the unmasked small molecule diffused through gap junctions. This effect can be
exploited to study cell-cell connectivity, but would confound cell-specific protein labeling if
the small molecule were to diffuse to cells lacking the decaging enzyme [
16
]. To address
these concerns, Tape
et al
. optimized CTAP for eukaryotic cell types and achieved ~90%
cell-specific labeling in ten-day co-cultures [
17
]. Using their optimized method, Tape
et al
.
combined CTAP with phosphoproteomics to study heterocellular KRAS
G12D
signaling in
pancreatic ductal adenocarcinoma cells [
18
]. By restricting their proteomic analysis to cells
that expressed KRAS
G12D
, the authors showed that the oncogene regulates AKT through
reciprocal signaling – not through the accepted cell-autonomous pathway.
Bio-orthogonal Non Canonical Amino acid Tagging (BONCAT)
CTAP is most suitable for cell-specific experiments conducted in culture on timescales of 3–
7 days [
19
]. For studies that require better time resolution, the bio-orthogonal non-canonical
amino acid tagging (BONCAT) method, introduced by Dieterich and coworkers, offers a
good alternative [
20
,
21
]. In its original form, BONCAT exploits the capacity of the
endogenous aminoacyl-tRNA synthetases to charge non-canonical amino acids (ncAAs) to
their cognate tRNAs for incorporation into proteins. ncAAs bearing bio-orthogonal chemical
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handles, often azides or alkynes, enable conjugation to affinity tags and separation of tagged
proteins from the rest of the protein pool. The methionine surrogates azidohomoalanine
(Aha) and homopropargylglycine (Hpg) have been used to probe proteome dynamics in
bacterial [
22
–
26
] and mammalian [
27
] systems, and notably, to enrich and quantify secreted
proteins [
28
]. Depletion of cellular methionine is not necessary for Aha labeling; Bagert
et
al
. showed that a 30:1 ratio of Aha to Met yielded excellent protein labeling while
minimizing perturbations that might be expected to arise from methionine starvation [
29
].
Other studies have shown that ncAA labeling for periods of up to two days do not perturb
embryonic growth in live mice [
30
]. In designing a BONCAT experiment, the investigator
should choose concentrations of the ncAA label and its natural counterpart that reflect the
relative rates of activation of the amino acids by the cognate synthetase.
In 2009, Ngo and coworkers developed a cell-selective version of BONCAT by engineering
an
E. coli
methionyl-tRNA synthetase (
Ec
MetRS) variant that activates azidonorleucine
(Anl). Because Anl is a poor substrate for wild-type
Ec
MetRS, labeling is essentially
restricted to cells that express the mutant synthetase. In the first example of the cell-specific
BONCAT method, Ngo
et al
. reported specific labeling of
E. coli
cells co-cultured with
murine alveolar macrophages [
31
]. Grammel
et al
. expanded on this method by enriching for
proteins synthesized during
Salmonella typhimurium
infection [
32
], and Mahdavi and
coworkers used BONCAT to determine the order in which
Yersinia enterocolitica
effector
proteins are injected into HeLa cells in the course of infection [
33
].
Cell-selective BONCAT has now been extended to proteomic analysis in live animals,
highlighting its potential utility in creating cell-specific proteomic “atlases”. In 2015 we
reported a mutant phenylalanyl-tRNA synthetase (PheRS) that enables the use of
p
-
azidophenylalanine (Azf) as a BONCAT probe in
Caenorhabditis elegans
[
34
]. Combining
cell-selective BONCAT with stable isotope labeling, we used the
myo-2
promoter to direct
expression of the mutant synthetase to the 20 pharyngeal muscle cells of the worm. We were
able to quantify 2270 proteins by this method, and to verify the pharyngeal expression
patterns of several previously uncharacterized proteins.
Dieterich and coworkers have adapted cell-selective BONCAT labeling to
Drosophila
melanogaster
through controlled expression of the
Dm
MetRS L262G mutant [
35
]. Chronic
administration of Anl in developing flies expressing the mutant synthetase caused slight
impairments in larval growth and behavior, but shorter (48 h) labeling times led to no
noticeable defects. Importantly, administration of the amino acid in flies that did not express
the mutant MetRS caused no discernible effect. Using this strategy, Niehues
et al
. measured
reduced neuronal protein synthesis rates in a
Drosophila
model of Charcot-Marie-Tooth
(CMT) neuropathy. Mahdavi
et al.
and Muller
et al.
have employed the analogous (L274G)
mouse synthetase in mammalian cell culture and in a neuronglia co-culture system,
respectively [
36
,
37
]. The latter experiments enabled the investigators to monitor changes in
the astrocytic proteome in response to treatment with brain-derived neurotrophic factor
(BDNF).
Split synthetases have been developed to enable cell-selective analysis of systems in which
no single promoter restricts expression of the mutant enzyme to the cells of interest [
38
].
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Notably, all amino acids and enrichment media needed for BONCAT experiments are
commercially available.
Stochastic Orthogonal Recoding of Translation (SORT)
Chin and coworkers have developed a residue-specific ncAA-labeling technology termed
stochastic orthogonal recoding of translation (SORT), which – like BONCAT – allows
chemoselective modification and enrichment of newly synthesized cellular proteins. SORT
relies on expression of a pyrrolysyl-tRNA synthetase and its cognate tRNA [
39
,
40
]. Using
this method, Elliott
et al
. cell-selectively labeled and identified proteins made during
different stages of larval growth in
Drosophila
. Importantly, SORT allows the anticodon of
the cognate tRNA to be changed to direct the ncAA to different sets of codons in the labeled
proteins. Elliott
et al
. have characterized the enrichment process and found that tagging at
different codons leads to the enrichment of overlapping, but distinct sets of proteins [
41
].
The authors noted that simultaneous expression of multiple tRNAs (i.e., tRNA-Ala, -Ser and
-Met) increases labeling efficiency. Furthermore, Elliott
et al.
found that enrichment after
tagging improves detection of low-abundance proteins.
Cell-selective O-propargyl-puromycin (OP-Puro) labeling
The
O
-propargyl-puromycin (OP-Puro) method also incorporates “clickable” handles into
nascent proteins [
42
]. Cohen and coworkers recently achieved cell-targeted OP-puromycin
labeling by using a phenylacetyl-caged analog that is uncaged by cell-selective expression of
penicillin G acylase (PGA) [
43
]. The OP-puro method is the fastest of the metabolic labeling
methods and the best suited for studies requiring ultra-short labeling times [
44
]. Prolonged
labeling with OP-puro would be expected to perturb cellular behavior through inhibition of
global translation. Furthermore, premature truncation renders this method ineffective for the
identification of secreted proteins.
Spatially restricted & subcellular proteomics
Ting and coworkers first used a mutant ascorbate peroxidase (APEX) to selectively tag
proteins localized to the mitochondrial matrix [
45
,
46
]. Unlike the cell-selective metabolic
labeling methods just described, this method labels all proteins, including pre-existing
proteins, within a subcellular volume. Chen
et al
. used this elegant strategy to characterize
multiple cell types in
Drosophila
, including the mitochondrial matrix of muscle tissue [
47
].
The Weissman laboratory has combined the APEX labeling method with ribosome profiling
to characterize localized protein synthesis in yeast [
48
,
49
]; extension of their method to cell-
selective analysis is readily imagined.
Choosing a cell-selective proteomic method
The choice of a cell-selective method of proteomic analysis should reflect careful
consideration of the advantages and disadvantages of each of the available approaches
(Table 1).
Physical sorting methods allow straightforward characterization of the steady-state proteome
of the cell type of interest. However, removing cells from their natural environments prior to
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analysis raises concerns about artifacts, leads to limited temporal information, and sacrifices
information about secreted proteins.
Ribosome profiling, when combined with cell-selective TRAP, provides significantly higher
coverage of the gene expression profile than any direct proteomic measurement. But
ribosome profiling is not a perfect proxy for protein synthesis and yields no information
regarding protein secretion [
50
]. Moreover, only direct proteomic methods allow detection
of post-translational modifications.
CTAP simplifies quantitative proteomic measurements for samples of relatively low
complexity, but enrichment-based strategies (i.e., BONCAT, SORT or OP-Puro) are likely to
be superior for short labeling times or for analysis of rare cells in complex tissues. Only
APEX yields snapshots of the steady-state proteome with sub-cellular resolution. All cell-
selective, enrichment-based experiments require the use of genetically tractable organisms.
Optimization of enrichment-based strategies requires careful consideration of alternative
purification chemistries. Attachment to the resin used for purification can be accomplished
either by direct covalent ligation or by a two-step process of affinity-tagging (e.g., with
biotin reagents) and non-covalent binding (e.g., to streptavidin resins). Following
appropriate washing steps, samples can be released from the resin by competitive binding,
by proteolysis, or by selective cleavage of the affinity reagent. APEX appends biotin to
surrounding molecules, so streptavidin-based resins are used to enrich for labeled proteins
[
46
]. OP-Puro requires an azide-based affinity handle or resin for enrichment [
43
]. SORT
uses cyclopropene labels and tetrazine linkers in a ligation reaction reported to be 100 to
1000 times faster than the strain-promoted azide-alkyne cycloaddition [
41
]. BONCAT labels
with either alkynes or azides, and enriches with complementary azide or alkyne reagents. A
special consideration arises in the analysis of lysates labeled with azides: Free thiols, which
are known to react with cyclooctynes, must be blocked with capping reagents such as
iodoacetamide or N-ethylmaleimide to avoid high background [
34
]. Many azide and alkyne
resins and linkers are commercially available, and tetrazine-based reagents are beginning to
appear on the market.
If the investigator wishes to identify the sites at which protein labeling has occurred, linkers
with cleavable moieties can be used [
51
]. For many experiments, though, identification of
labeling sites is not necessary, and on-bead digestion of enriched proteins is often simpler
and more straightforward. In our hands, directly conjugating azide-labeled lysates to
cyclooctyne resins has allowed us to identify larger numbers of relevant proteins [
34
].
Because enrichments are never perfect, running mock enrichments of unlabeled sample
along with labeled samples provides a useful indication of background reactivity and non-
specific protein contamination. Samples with abundant contaminating biopolymers such as
pectin, serum proteins, or mucin may need an additional step to remove or degrade these
contaminants and facilitate successful enrichment.
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Conclusions and future outlook
Recent years have witnessed the introduction of powerful techniques that allow investigators
to monitor protein synthesis with unprecedented resolution in space and time. Cell-specific
proteomic analyses will play a key role in the identification of the mechanisms that govern
cell specialization and that allow complex organisms to respond to changing environments.
Acknowledgments
Funding:
Caltech research on cell-specific proteomic analysis has been supported by NIH grants R01-GM062523
and R21-AI121890, and by the Institute for Collaborative Biotechnologies through grant W911NF-09-0001 from
the U.S. Army Research Office.
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S, Bunse S, et al. Direct visualization of newly synthesized target proteins in situ. Nat Meth. 2015;
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Highlights
•
Cell-selective proteomics is important in complex, heterocellular
environments
•
Innovative chemical tools enable unbiased cell-type-specific interrogation of
translation
•
Labeling methods including TRAP, CTAP, BONCAT, SORT, OP-Puro and
APEX have been developed for cell-selective analysis
•
Sequencing and mass spectrometry-based strategies complement one other in
the study of protein synthesis
•
The strengths and limitations of each analytical method must be considered
carefully in the context of the biological question to be addressed
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Figure 1.
The importance of cell-type-specific proteomics. Bulk measurements of complex tissues can
obscure proteomic changes that occur in specific sub-populations of cells. A protein that is
highly expressed (up arrows) in the cells of interest might be detected at low abundance
overall due to low expression (down arrows) in background cells. Cells of interest must be
physically isolated or tagged to measure the cell-specific proteome. Physical isolation
measures steady-state levels of intracellular proteins, whereas labeling methods can be time-
resolved and used to identify secreted proteins.
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Figure 2.
Labeling strategies for cell-selective proteomics. a) The process by which amino acids are
incorporated into proteins, and the step exploited by each of the labeling methods discussed
in this Opinion. b) Schematic of each technique.
Translating ribosome affinity purification: TRAP; Cell type-specific labeling using amino
acid precursors: CTAP; Bio-orthogonal non-canonical amino acid tagging: BONCAT;
Stochastic orthogonal recoding of translation: SORT;
O
-propargyl puromycin: OP-Puro;
ascorbate peroxidase: APEX; Lysine racemase:Lyr; diaminopimelate decarboxylase: DDC;
aminoacyl-tRNA synthetase: RS; penicillin-G-acylase: PGA.
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Table
Advantages and disadvantages of cell-specific proteomic methods.
Cell-
Specific
Method
Biomolecule
identified
Organisms
demonstrated
in?
Temporal Resolution
Secreted?
PTM?
Advantages
Disadvantages
References
Translatomics
TRAP
mRNA
Prokaryotes,
eukaryotes
Snapshot of translation
No
No
High sequence coverage,
able to combine with
ribosome profiling
Requires expression of tagged
ribosome, miss
translational control
[
7
–
11
]
Cell
Separation
Manual
mRNA, Protein
Prokaryotes,
eukaryotes,
clinical samples
Steady-state proteome
No
Yes
Straightforward, inexpensive
Possible artifacts from sample
preparation,
time and labor intensive
[
3
], [
12
–
13
]
FACS
mRNA, Protein
Prokaryotes,
eukaryotes,
clinical samples
Steady-state proteome
No
Yes
High-throughput
Requires dissociation of cells,
need
expression of transgene or
recognizable
epitope, need specialized
equipment
[
14
]
Metabolic
CTAP
Protein
Cell culture
Up to 10 days
continuous cell culture
Yes
Yes
Quantitative, compatible with
long-term cell culture
Requires expression of Lyr/
DDC, cells must
be auxotrophic for lysine,
restricted to cell
culture
[
15
], [
17
–
19
]
BONCAT
Protein
Prokaryotes,
eukaryotes, cell
culture
Newly synthesized
proteins in minutes
(prokaryotic, cell
culture) to days (whole
animal)
Yes
Yes
Commercially available
reagents, high degree of
temporal resolution
Requires expression of
synthetase; only
Met/Phe residue replacement
available
currently, requires delivery of
the ncAA
[
20
–
38
], [
44
]
SORT
Protein
Eukaryotes, cell
culture
Newly synthesized
proteins in minutes (cell
culture) to days (whole
animal)
Yes
Yes
Easy to change the residue
of non-canonical amino acid
incorporation, high degree of
temporal resolution
Requires expression of
synthetase/tRNA pair,
reagents not currently
commercially available,
requires delivery of the ncAA
[
39
–
41
]
OP-Puro
Protein
Cell culture
Newly synthesized
proteins in minutes (cell
culture)
No
No
Not residue-dependent,
highest degree of temporal
resolution
Requires expression of PGA,
reagents not
currently commercially
available
[
42
–
44
]
Spatial
APEX
mRNA, Protein
Eukaryotes, cell
culture
Subcellular, steady-
stateproteome
No
Yes
High degree of spatial
resolution (subcellular)
Requires expression of APEX/
HRP, no
temporal resolution, requires
delivery of
peroxide + biotin-phenol
reagent, possible
toxicity of peroxide over longer
timescales,
[
45
–
49
]
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. Author manuscript; available in PMC 2018 February 01.