Biocompatible click chemistry enabled compartment-specific pH measurement inside E. coli

Abstract

Bioorthogonal reactions, especially the Cu(I)-catalysed azide-alkyne cycloaddition, have revolutionized our ability to label and manipulate biomolecules under living conditions. The cytotoxicity of Cu(I) ions, however, has hindered the application of this reaction in the internal space of living cells. By systematically surveying a panel of Cu(I)-stabilizing ligands in promoting protein labelling within the cytoplasm of Escherichia coli, we identify a highly efficient and biocompatible catalyst for intracellular modification of proteins by azide-alkyne cycloaddition. This reaction permits us to conjugate an environment-sensitive fluorophore site specifically onto HdeA, an acid-stress chaperone that adopts pH-dependent conformational changes, in both the periplasm and cytoplasm of E. coli. The resulting protein-fluorophore hybrid pH indicators enable compartment-specific pH measurement to determine the pH gradient across the E. coli cytoplasmic membrane. This construct also allows the measurement of E. coli transmembrane potential, and the determination of the proton motive force across its inner membrane under normal and acid-stress conditions.

ARTICLE

Received 1 Jun 2014 | Accepted 13 Aug 2014 | Published 19 Sep 2014

Bioorthogonal reactions, especially the Cu(I)-catalysed azidealkyne cycloaddition, have revolutionized our ability to label and manipulate biomolecules under living conditions. The cytotoxicity of Cu(I) ions, however, has hindered the application of this reaction in the internal space of living cells. By systematically surveying a panel of Cu(I)-stabilizing ligands in promoting protein labelling within the cytoplasm of Escherichia coli, we identify a highly efcient and biocompatible catalyst for intracellular modication of proteins by azidealkyne cycloaddition. This reaction permits us to conjugate an environment-sensitive uorophore site specically onto HdeA, an acid-stress chaperone that adopts pH-dependent conformational changes, in both the periplasm and cytoplasm of E. coli. The resulting protein uorophore hybrid pH indicators enable compartment-specic pH measurement to determine the pH gradient across the E. coli cytoplasmic membrane. This construct also allows the measurement of E. coli transmembrane potential, and the determination of the proton motive force across its inner membrane under normal and acid-stress conditions.

1 Synthetic and Functional Biomolecules Center, Beijing National Laboratory for Molecular Sciences, Key Laboratory of Bioorganic Chemistry and Molecular Engineering of Ministry of Education, College of Chemistry and Molecular Engineering, Peking University, Beijing 100871, China. 2 Department of Biochemistry, Albert Einstein College of Medicine of Yeshiva University, 1300 Morris Park Avenue, Bronx, New York 10461, USA. 3 State Key Laboratory of Pharmaceutical Biotechnology, School of Life Sciences, Institute of Chemistry and BioMedical Sciences, Nanjing University, Nanjing 210093, China. 4 Peking-Tsinghua Center for Life Sciences, Peking University, Beijing 100871, China. Correspondence and requests for materials should be addressed to J.Z. (email: mailto:[email protected]

Web End [email protected] ) or to P.W. (email: mailto:[email protected]

Web End [email protected] ) or to P.R.C. (email: mailto:[email protected]

Web End [email protected] ).

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DOI: 10.1038/ncomms5981

Biocompatible click chemistry enabled compartment-specic pH measurement inside E. coli

Maiyun Yang1, Abubakar S. Jalloh2, Wei Wei3, Jing Zhao3, Peng Wu2 & Peng R. Chen1,4

ARTICLE NATURE COMMUNICATIONS | DOI: 10.1038/ncomms5981

Both eukaryotic and prokaryotic cells are compartmentalized. Inside eukaryotic cells, metabolic processes and signalling events are frequently carried out in the cytosol

or specialized organelles (for example, mitochondria, ER and Golgi), with well-dened pH and oxidative status. Although to a lesser extent, the intracellular space of Gram-negative bacteria is also compartmentalized as cytoplasmic and periplasmic spaces, with the latter separated from the environment and the cytoplasm by a highly porous outer membrane and a tighter inner membrane (or cytoplasmic membrane), respectively. This arrangement produces a distinct environment within these bacterial compartments under normal and stress conditions. For example, enteric pathogens such as Escherichia coli and Shigella spp. have to pass through the highly acidic human stomach (pHo3) before reaching their primary infection site in the small intestine1,2. To survive this acidic environment, E. coli cells have evolved multiple acid-resistance systems to elevate their internal pH3, including generating a pH gradient across the cytoplasmic membrane. The pH gradient (DpH

pHcytoplasm pH

periplasm) is a key component of the proton motive force (PMF), which, in conjunction with the membrane potential (DC), determines the electrochemical gradient, namely

PMF, across E. coli cytoplasmic membrane4. Many biological processes are energetically linked to the free energy produced by PMF, including ATP synthesis, the transport of nutrients across the cytoplasmic membrane, as well as the rotation of bacterial agella5,6. There are currently no suitable indicators for measuring pH gradient under acid-stress, since small molecule uorophores lack targeting specicity while pH-sensitive uorescent proteins denature below pH5 (refs 7,8). Therefore, the ability to directly target pH indicators into different E. coli compartments is highly desired.

Coupling the genetic code expansion strategy with bioorthogonal chemistry provides a powerful tool for highly specic protein labelling in vitro and in living cells. For example, an unnatural amino acid bearing a bioorthogonal handle can be genetically incorporated into a given protein that is expressed in a specic location, allowing the subsequent bioorthognal labelling with a small molecule uorophore. However, this strategy has largely focused on in situ labelling of biomolecules topologically located on the surface of mammalian or bacterial cells9,10, or within the bacterial periplasm11,12.

Protected by single or double plasma membranes, molecules located in the highly reduced and fragile cytoplasm represent attractive yet challenging targets for bioorthogonal labelling. Currently, the state-of-the-art bioorthogonal click reactions include the Cu(I)-catalysed azidealkyne cycloaddition (CuAAC) and the strain-promoted azidealkyne cycloaddition (SPAAC), among a few others1315. In their pioneering work on SPAAC, Tirrell, Bertozzi and co-workers found that, when cyclooctyne-based uorescent probes was used to label newly synthesized proteins in live mammalian cells16, a high uorescence background was observed, which was later attributed to the nonspecic reactivity of the DIFO probe toward free thiols or cysteine-containing proteins17,18. Notably, several studies have shown that CuAAC exhibited 10 100 times faster kinetics than

SPAAC in aqueous solutions, and that the terminal alkyne is an excellent bioorthogonal handle19,20. These attributes make CuAAC an attractive candidate for in vivo labelling.

However, copper is known to be toxic to both eukaryotic and prokaryotic cells. For example, copper destroys many biomolecules by oxidative damage, and thus, E. coli compartmentalizes its copper-dependent enzymes in the periplasm as well as the outer aspect of the cytoplasmic membrane, leaving an extremely low level of copper in the reduced cytoplasm21. Furthermore, several recent studies showed that the highly thiophilic Cu(I) ions can

directly impair Fe-S cluster-containing enzymes located exclusively within the bacterial cytoplasm, which has been suggested as a major lethal effect of copper inside microorganisms22,23. Interestingly, these same studies indicated that sequestration of copper ions by chelators such as bathocuproine sulphonate or copper-binding proteins can restrict the tendency of copper to damage intracellular Fe-S clusters, and thus enhance bacterial tolerance to copper. These observations, together with our recent success in the discovery of accelerating ligands that render CuAAC biocompatible for labelling cell-surface glycans in living organisms24,25, prompted us to explore the feasibility of utilizing the ligand-assisted CuAAC to label cytoplasmic proteins within living bacterial cells.

Herein, we report that tris(triazolylmethyl)amine-coordinated Cu(I) catalysts, BTTPCu(I) and BTTAACu(I), permit the in situ labelling of azide-incorporated proteins in the cytoplasm of E. coli without apparent toxicity. Employing this biocompatible ligation chemistry, we specically targeted a proteinuorophore hybrid pH indicator into the E. coli cytoplasm for internal pH measurement. By employing both the cytoplasm- and periplasm-residing pH indicators, we determine the pH values in these two compartments under highly acidic conditions. The calculated pH gradient (DpH) across E. coli cytoplasmic membrane, in conjunction with the measured transmembrane potential (DC) using a DC-sensitive dye, enable us to obtain the PMF value across E. coli cytoplasmic membrane under acid-stress conditions.

ResultsLigand-assisted CuAAC for protein labelling in bacterial cytoplasm. As the rst step to evaluate CuAAC as a biocompatible tool to label cytosolic proteins in E. coli, we chose a cytosolically expressed green uorescent protein (GFP) bearing a single azide handle as the model system. A panel of Cu(I)-stabilizing ligands developed by us and others were surveyed to assess their efciency in promoting the CuAAC-mediated protein labelling (Fig. 1a). TBTA, the canonical ligand developed by Sharpless and co-workers, is the most commonly used ligand for bioorthogonal conjugation26. However, TBTA has poor solubility in aqueous buffer, resulting in incomplete ligation when azide and alkyne are at micromolar concentrations27. BTTAA and BTTPS are two water soluble TBTA analogues. Both ligands dramatically accelerate CuAAC and render it biocompatible for labelling cell-surface glycans and proteins in live zebrash embryos24,25. As an unsulphated version of BTTPS, BTTP exhibited a similar efciency as BTTPS in promoting CuAAC-mediated protein in vitro labelling25. Finally, we also examined bathophenanthroline disulphonate (BPS), a well-known negatively charged Cu(I)-chelator, as well as L-histidine, a recently reported effective ligand for accelerating CuAAC to label mammalian cell-surface glycans28,29.

The azide-bearing GFP (GFP-N149-ACPK), constructed by site-specic incorporation of a pyrrolysine (Pyl) analogue bearing an azide residue, ACPK (Fig. 1a), into residue 149 via our previously evolved pyrrolysyl-tRNA synthetase-tRNA pair, was expressed in E. coli. We performed the ligation chemistry by incubating live E. coli cells harbouring GFP-N149-ACPK with alk-4-DMN in the presence of the aforementioned panel of copper ligands at room temperature for 1 h (Fig. 1b, Supplementary Figs 14). The lysates of cells were resolved by SDSpolyacrylamide gel electrophoresis (SDSPAGE) and analysed by in-gel uorescence. As shown in Fig. 1b, BTTPCu(I) and BTTAACu(I) exhibited the highest reactivity in catalyzing this in vivo protein labelling process; the yield of BTTPCu(I)-mediated reaction was at least sevenfold higher than that achieved by using the uncoordinated Cu(I) (Supplementary Fig. 5).

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NATURE COMMUNICATIONS | DOI: 10.1038/ncomms5981 ARTICLE

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Figure 1 | Comparison of the labelling efciency in vitro and within the E. coli cytoplasm. (a) Structures of the Pyl analogue ACPK, the alkyne-tethered uorophore alk-4-DMN as well as Cu(I)-stabilizing ligands for CuAAC reaction used in this study. ACPK, Ne-((1R,2R)-2-azidocyclopentyloxy)carbonyl)-L-lysine; BPS, bathophenanthroline disulphonate disodium salt; BTTAA, 2-[4-{(bis[(1- tert-butyl-1H-1,2,3-triazol-4-yl)methyl]amino)methyl}-1H-1,2,3-triazol-1-yl]-acetic acid; BTTP, 3-[4-({bis[(1-tert-butyl-1H-1,2,3-triazol-4-yl)methyl]amino}methyl)- 1H-1,2,3-triazol-1-yl]propanol; BTTPS, 3-[4-({bis[(1-tert-butyl-1H-1,2,3-triazol-4- yl)methyl]amino}methyl)-1H-1,2,3-triazol-1-yl]propyl hydrogen sulphate; 4-DMN: 4-N,N-dimethyl amino-1,8-naphthalimide; TBTA, tris-((1-benzyl-1H-1,2,3-triazol-4-yl)methyl)amine. (b,c) Efciency of in vivo (b) and in vitro (c) protein labelling mediated by ligand-assisted CuAAC. The reaction products between GFP-N149-ACPK and alk-4-DMN were analysed by SDSPAGE and visualized under ultraviolet illumination (top) before being stained by Coomassie blue (bottom). Blank, without Cu(I) catalyst; M, protein marker; MFI, mean uorescence intensity.

Approximately two- to threefold lower activity was observed in the TBTACu(I) and L-histidine-Cu(I) catalysed reactions. By contrast, BTTPSCu(I) yielded an extremely low amount of uorescently labelled proteins and no detectable uorescent product was obtained in the BPSCu(I) mediated reaction. These results were in direct contrast to the data obtained from the in vitro labelling experiments (Fig. 1c), in which equal quantity of GFP-N149-ACPK was reacted with alk-4-DMN in the presence of the same set of Cu(I) complexes. In the in vitro labelling experiments, BPSCu(I) and BTTPSCu(I) produced similar levels of ligation products compared with that of BTTPCu(I) and BTTAACu(I), whereas TBTA and L-histidine exhibited two-

to threefold lower labelling efciency than the remaining four ligands. In addition, the GFP protein still retained about 95% of its uorescence intensity after the treatment with BTTPCu(I) and BTTAACu(I), indicating that these Cu(I) complexes had little inuence on its structural integrity (Supplementary Fig. 6).

The difference between the efciency of Cu(I) catalyst-assisted click labelling of GFP-N149-ACPK in vitro and within livingE. coli cells suggests that even though the same concentrations of ligandCu(I) complexes were used, their working concentrations in bacterial cytoplasm might be signicantly varied. To test this hypothesis, we employed ICPAES (inductively coupled plasmaatomic emission spectrometry) to measure the copper

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ARTICLE NATURE COMMUNICATIONS | DOI: 10.1038/ncomms5981

concentration within E. coli cytoplasm upon treatment with these Cu(I) complexes (Supplementary Methods and Fig. 2a). We isolated the cytoplasmic fraction of bacteria using an osmotic shock method to avoid contamination by copper from the periplasm (Supplementary Fig. 7). ICPAES analysis on these cytoplasmic samples and whole-cell samples showed that the bacterial cells exhibited apparent preference towards certain copper sources. More copper was present in E. coli cytoplasm when treated with free Cu(I) ions than with an equal amount of ligandCu(I) complexes. As expected, BPSCu(I) and BTTPS Cu(I) were taken up by the cells less effectively compared with other Cu(I) complexes because the negatively charged sulphate group on BPS and BTTPS signicantly decreased their membrane permeability. This observation, combined with the aforementioned in vivo labelling results, suggest that the low labelling efciency observed in the BPSCu(I) and BTTPSCu(I) catalysed reactions was likely due to the low level of these Cu(I) complexes delivered into the cytoplasm. Importantly, although the other four Cu(I) complexes resulted in equimolar Cu(I) concentration within the cytoplasm, their ability to promote the CuAAC labelling reaction varied signicantly, with BTTP and BTTAA possessing two- to threefold higher efciency than the other two ligands. Taken together, our results indicated that, at equal concentration of Cu(I), BTTP and BTTAA served as the best ligands for accelerating CuAAC reaction in the bacterial cytoplasm.

Toxicity study of Cu(I) complexes to E. coli. Next, we investigated the toxicity of these Cu(I) complexes to E. coli cells by using a proliferation assay. As shown in Fig. 2b, similar to the uncoordinated Cu (I) ions, the TBTACu(I) complex dramatically inhibited bacterial proliferation. By contrast, all the other ligands attenuated copper toxicity to a certain extent. In particular, BTTPSCu(I) and BTTPCu(I) imparted negligible inuences on bacterial growth. To evaluate the potential harmful effects from

these Cu(I) complexes after a longer incubation time, a plate sensitivity assay was also performed on bacterial cells treated with the same panel of Cu(I) copper complexes overnight, which yielded similar results (Supplementary Fig. 8). The low toxicity of BTTPS might be due to its low membrane permeability whereas BTTP and BTTAA might effectively block the damaging effects of Cu(I) to essential intracellular proteins such as Fe-S cluster-containing enzymes. To determine if this ligand coordination attenuates the damage of Fe-S cluster-containing enzymes by Cu(I) ions, we adopted a cell-growth assay using E. coli GR17 strain that lacks copper homeostatic systems (copA::kan DcueO DcusCFBA::cm) developed by Imlay and co-workers22 (Supplementary Fig. 9). Treatment with 10 mM of uncoordinated Cu(I) was sufcient to inhibit the growth of GR17 cells, whereas inclusion of the ligands BTTP or BTTAA restored cell growth almost to the same level as the addition of branched-chain amino acids. These results further conrmed that BTTP and BTTAA could largely sequester Cu(I) induced damage of Fe-S cluster-containing enzymes such as isopropylmalate dehydratase involved in the biosynthesis of branched-chain amino acids.

The mechanism of copper-associated cytotoxicity remains elusive. A long-standing hypothesis is that Cu(II)/Cu(I) ion redox chemistry mediates the production of reactive oxygen species (ROS)30 such as the highly deleterious hydroxyl radicals, which may cause damages to the cell membrane and Fe-S cluster-containing enzymes22. We used a commercial uorogenic assay to assess if ligand coordination attenuates the production of hydroxyl radicals generated by Cu(I)31 (Supplementary Fig. 10). All ligands examined reduced ROS production to a certain degree, with BTTPS and BTTP exhibiting the highest efciency. Furthermore, we directly analysed the Cu(I)-mediated ROS production inside E. coli cells by using a hydroxyl radical-sensitive uorescent probe 20,70-dichlorouorescein (DCF) as the reporter32 (Fig. 2c). Whereas the ligand-free Cu(I) ions generated a sevenfold increase in intracellular ROS compared with untreated cells, TBTACu(I) showed only a threefold

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Figure 2 | Toxicity of Cu(I) complexes inside E. coli cells. (a) Metal uptake of Cu(I)ligand complexes in the E. coli cytoplasm. Both the whole-cell fraction (blue columns) and the cytoplasmic fraction (red columns) were subjected to copper content measurement by ICPAES. Error bars, s.d. from three independent experiments. (b) E. coli growth curves after being treated with different Cu(I)ligands. Blank, E. coli cells without Cu(I) treatment. The data are representative of three independent experiments. (c) Detection of intracellular generation of ROS from E. coli cells using DCFH-DA as the reporter. Blank,E. coli cells without Cu(I) treatment. Error bars, s.d. from three independent experiments. (d) Fluorescence of PI-stained E. coli cells after being treated with Cu(I) ligands for 1 h. Blank, E. coli cells without Cu(I) treatment. Error bars, s.d. from three independent experiments.

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uorescence increase and all other Cu(I) complexes exhibited a o1.5-fold uorescence change.

Effect of Cu(I) complexes on membrane integrity. Maintaining cell membrane integrity is critical for bacterial physiology. Accordingly, it is important to develop an accurate measurement of pH gradient and PMF values across the E. coli membrane. To ascertain the integrity of E. coli membrane upon treatment with Cu(I) complexes, we incubated the bacteria with each of the six Cu(I) catalysts ([Cu] 100 mM, [ligand] 200 mM) for 1 h at

room temperature, followed by propidium iodide (PI) staining33 (Fig. 2d). Severe membrane damage was observed when cells were treated with either uncoordinated Cu(I) or Cu(I)TBTA complex. The remaining Cu(I) ligands all effectively attenuated coppers damage to the membrane. Similar results were obtained when trypan blue34, another commonly used uorescent dye to assess cell viability, was used (Supplementary Methods and Supplementary Fig. 11). The propensity for ROS production by these Cu(I) catalysts correlates well with their membrane damaging effects.

Our in vitro and in vivo experiments demonstrate that uncoordinated Cu(I) was highly active in generating detrimental hydroxyl radicals, whereas ligand-coordinated Cu(I) complexes produced much lower levels of oxidative species and thus less toxic. This is likely because the ligand may actively adjust the redox potential of Cu(I) ions through coordination. In theory, the higher the redox potential, the lower the propensity of Cu(I) complexes in generating ROS35. To examine whether our aforementioned ndings are consistent with this principle, we measured the redox potentials of BTTPSCu(I), BTTPCu(I) and BTTAACu(I) complexes and compared our results with the well-documented redox potential of the uncoordinated Cu(I). As shown in Table 1 and Supplementary Fig. 12, the measured redox potentials of BTTPSCu(I) and BTTPCu(I) were approximately 40 mV higher than that of BTTAACu(I), which, in turn, was 90 mV higher than the value of the uncoordinated Cu(I).

Taken together, our experiments conrmed that the ligand-free Cu(I) and TBTA-complexed Cu(I) were highly toxic to E. coli cells and that BPS- and BTTPS-complexed Cu(I) exhibited low toxicity, likely due to the negatively charged sulphate group that blocks their cellular entry. However, this same feature also rendered these two catalysts unsuitable for intracellular applications. Consistent with the previous study in mammalian cells,

L-histidine was found to attenuate copper-associated toxicity inE. coli. However, its ability to accelerate CuAAC is considerably lower than the water soluble tris(triazolylmethyl)amine-based ligands. In addition, there are concerns that L-histidine may serve as a solubilizing agent for Cu(I) to facilitate damaging of essential Fe-S clusters of dehydratases22. By increasing the redox potential of the coordinated Cu(I), BTTP and BTTAA signicantly reduced the production of ROS as well as the damaging effects to cell membrane. These ligands also efciently blocked the damage of Cu(I) to intracellular Fe-S cluster containing enzymes. Together, excellent biocompatibility inside living bacterial cells can be

achieved with BTTP and BTTAA. In particular, when compared with the BTTAACu(I) complex that we have previously used12, we found that BTTPCu(I) exhibited a further lowered ROS production, cellular damage and toxicity in assisting CuAAC-mediated protein labelling inside living E. coli cells. This sets up the stage for applying BTTPCu(I) complex for protein labelling within internal spaces of live bacterial cells.

Retargeting a periplasmic pH indicator into the cytosol. To demonstrate the applications of BTTP-assisted CuAAC for intracellular protein labelling beyond GFP, we applied this bio-compatible ligation chemistry to target a previously developed, periplasm-located protein-small molecule pH indicator into theE. coli cytoplasm. Enteric pathogens such as E. coli have to pass through the highly acidic human stomach (pHo3) before reaching their primary infection site in the small intestine. Due to the highly porous nature of the bacterial outer membrane, the periplasmic space is believed to rapidly equilibrate with the acidic extracellular environment7,36. In contrast, the cytoplasm is buffered at a much higher pH level, thereby generating an intracellular pH gradient that is crucial for the survival of enteric bacteria during acid-stress.

Compartment-specic pH indicators derived from the same uorphore or uorescent protein is advantageous for quantitative comparison of pH values within different intracellular spaces, mainly due to their unied chemical nature and pH response features. However, in contrast to many organelle-specic derivatives of a pH indicator that is applicable to different mammalian compartments, it remains a challenge to directly target a unied indicator into different bacterial spaces for pH measurement. The pH-responsive uorescent proteins are largely restricted to the E. coli periplasm due to the difculty for cytoplasmic-membrane trafcking, as well as the low efciency in folding of these uorescent proteins in the oxidized periplasm. Furthermore, most of these pH-sensitive proteins will be denatured when the pH drops to o5, rendering them incapable of measuring highly acidic environment that resembles those met in the human stomach. In addition, most small molecule uorophores lack targeting specicity to discriminate between the bacterial periplasm and cytoplasm. The membrane permeable uorophores may occupy both spaces, whereas bulky or negatively charged uorophores are inaccessible to the cytoplasm due to the tight cytoplasmic inner-membrane space. Therefore, the lack of a unied indicator suitable for pH measurement in different internal bacterial spaces renders the internal pH gradient of E. coli cells rather elusive, particularly under acid-stress conditions.

We recently developed a periplasm-localizing, protein uorophore hybrid pH indicator by applying BTTAA-assisted CuAAC between an alkyne-bearing solvatochromic uorophore (alk-4-DMN) and a periplasm-expressing acid chaperone HdeA, with ACPK incorporated as residue 58 within its pH-responsive region12 (peri-HdeA-58-ACPK; Fig. 3a). The resulting periplasm-residing pH indicator, termed peri-pHin, was able to operate under a wide pH range, permitting us to specically measure the acidity of the E. coli periplasm when the extracellular pH varied from neutral to pH 2. Because removal of the amino (N)-terminal signalling peptide can redirect HdeA from the periplasm to the cytoplasm12 and our BTTPCu(I) complex is highly compatible with the cytoplasm, we employed BTTP-assisted CuAAC to label the cytosolic version of ACPK-bearing HdeA protein (cyto-HdeA-58-ACPK) with alk-4-DMN. The proper localization of the resulting cytosolic pH indicator, termed cyto-pHin, was veried by immunoblotting analysis against the cytosolic as opposed to the periplasmic fraction of E. coli using an anti-HdeA antibody

Table 1 | Electrochemical data of Cu(I) complex in H2O.

Cu(I) complexes E1/2(mV) DEp (mV) BTTPCu(I) 50 232

BTTPSCu(I) 49 251 BTTAACu(I) 42 191 Cu(I) 30 101

E 1/2 (Ea Ec), DEp |Ea Ec|. Ea, Ec are dened as the potentials of the maximum of

current intensity when scanning toward anodic or cathodic potentials.

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a

peri-HdeA

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Figure 3 | Directing a genetically encoded click-labelled pH indicator to different E. coli internal spaces. (a) An acid chaperone protein HdeA with or without a periplasm-targeting signal peptide (SP) were expressed carrying a site-specically incorporated azide-bearing unnatural amino acid, ACPK, as residue 58. The resulting proteins were then conjugated with alk-4-DMN via ligand-assisted CuAAC reaction to afford specic pH indicators in E. coli cytoplasm (cyto-pHin) and periplasm (peri-pHin), respectively. IM, inner membrane (cytoplasmic membrane), OM, outer membrane, SP, signal peptide sequence. CuAAC: copper catalysed azidealkyne cycloaddition. (b) Immunoblotting analysis showing the localization of peri-HdeA and cyto-HdeA. (c) Generation of compartment-specic pH indicators, peri-pHin and cyto-pHin, as analysed by SDSPAGE. M, protein marker. (d) Flow cytometry results of E. coli cells harbouring cyto-pHin as a function of pH (7.02.0).

(Fig. 3b; Supplementary Fig. 13). Furthermore, the BTTP-assisted click-labelling efciency and the specicity for producing both peri-pHin and cyto-pHin were conrmed by uorometric SDS PAGE analysis (Fig. 3c). In vitro uorescence measurement showed that peri-pHin and cyto-pHin proteins have nearly the same pH response pattern (Supplementary Fig. 14). Notably, by quantitatively comparing the protein expression levels and the corresponding uorescence intensity from the two gel bands, the in vivo click-labelling efciency of cyto-pHin was determined to be 0.94 as that of peri-pHin (Supplementary Fig. 15). In addition, by quantitative comparison of the uorescence intensity between peri-HdeA58-ACPK (or cyto-HdeA58-ACPK) proteins labelled by alk-4-DMN in E. coli cells and the in vitro quantitatively labelled HdeA58-DMN, we have calculated the in vivo labelling yield as 73 and 69% for peri-HdeA58-ACPK and cyto-HdeA58-ACPK, respectively (Supplementary Fig. 16).

We next applied ow cytometry to measure the uorescence ofE. coli cells harbouring cyto-pHin under a wide range of extracellular acidity. The extracellular pH values were reduced step-wise from pH 7 to pH 2 followed by ow cytometric analysis

at each pH value (Fig. 3d). A gradual increase in uorescence signal was observed, and we plotted this uorescence change against each pH unit. A total of threefold uorescence enhancement in cyto-pHin signal was observed, which was smaller than that of peri-pHin (4vefold), indicating that the cytosolic pH varied to a lesser extent than the periplasmic pH.

Measuring pH gradient across the cytoplasmic membrane. By employing our two compartment-specic pH indicators, peripHin and cyto-pHin, we next measured the pH gradient acrossE. coli cytoplasmic membrane under acid-stress conditions. Flow cytometric analysis was rst performed on the peri-pHin-expressing cells under various external pH conditions with and without 20 mM benzoate, a membrane permeable weak acid that transports protons across the bacterial membranes to lower the internal pH37. Consistent with previous reports, our results revealed that the E. coli periplasmic pH showed negligible change in response to benzoate treatment when the extracellular pH was 45 (ref. 7). However, when the external pH dropped to o5, peripHin detected a small but noticeable change in uorescence after the benzoate treatment; the presence of benzoate led to a further increase in uorescence of peri-pHin (Fig. 4a). As a negative control, E. coli cells bearing the pH-insensitive variant peri-HdeA-72-DMN were subjected to ow cytometric analysis under different pH conditions, but exhibited essentially no uorescence changes throughout all pH conditions in the presence and absence of 20 mM benzoate (Supplementary Fig. 17). This led us to believe that the periplasmic space of E. coli might also be slightly buffered by basic amino acids and polyamines that are exported from the E. coli cytoplasm, particularly when surrounded by highly acidic environment such as gastric acid.

Next, we utilized E. coli cells harbouring cyto-pHin to monitor the cytoplasmic pH change in response to an increasing extracellular acidity from pH 7 to pH 2 with and without 20 mM benzoate (Fig. 4b). In contrast to the pH measurement in the periplasm, our analysis showed that the addition of benzoate enhanced the uorescence under all pH conditions, consistent with the fact that the E. coli cytoplasm maintains a much higher buffering capacity than its periplasm. In particular, the bacterial uorescence in the presence of benzoate increased signicantly when the extracellular acidity was decreased to below pH 5, which indicated a stronger cytosolic buffering capacity upon the acidication of the extracellular environment. We also expressed the pH-insensitive HdeA control variant (HdeA-72-DMN) in the cytoplasm (cyto-HdeA-72-DMN), but observed no noticeable uorescence differences under all pH conditions we tested with and without benzoate (Supplementary Fig. 17). For quantitative measurement, we generated a calibration curve of the extra-cellular pH as a function of relative uorescence intensity (RFI, Fig. 4c); the curve was further validated by a series of standard pH solutions (Supplementary Fig. 18).

Based on these pH-dependent ow cytometric investigations as well as the RFI derived from peri-pHin and cyto-pHin at a given pH buffer, we determined that the periplasmic pH was about 0.3 pH unit higher than the extracellular space when the environmental acidity was at pH 3.0. Under the same condition, E. coli cytosolic pH was calculated to be 4.4 (Fig. 4c,d), which is signicantly higher than the environment or the cytoplasm after benzoate treatment (Supplementary Fig. 19). Thus the pH gradient across the E. coli cytoplasmic membrane was about

1.1 pH unit when the extracellular pH was at 3.

To further verify the reliability of our pH indicators, we incubated E. coli cells harbouring the cyto-peri-pH indicator in acidic M9 minimal medium (pH 3) or in an acidic buffer containing pepsin, a digestive enzyme that is present in the gastric

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55

55

peri-pHin + benzoate peri-pHin benzoate

cyto-pHin + benzoate cyto-pHin - benzoate

Fluorescence intensity (a.u.)

Fluorescence intensity (a.u.)

45

45

35

35

25

25

15

15

5

5 7.0 6.0 5.0 4.0 3.0 2.0 7.0 6.0 5.0 4.0 3.0 2.0 pH pH

3.0

50

2.5

45

40

2.0

35

RFI

1.86

1.04

RFI

1.5

30

25

24.1

1.0

20

0.5

7.0 6.0 5.0 4.0 3.0 2.0

150

120

20 40 60 80 100

160

140

pH

[afii9820] ( mV )

Figure 4 | Measuring the pH values in E. coli periplasm and cytoplasm under acid stress. (a) The pH-dependent uorescence response curves of peri-pHin in the presence and absence of 20 mM sodium benzoate. Error bars, s.d. from three independent experiments. (b) The pH-dependent uorescence response curves of cyto-pHin in the presence and absence of 20 mM sodium benzoate. Error bars, s.d. from three independent experiments. (c) Calibration curve of extracellular pH as a function of relative uorescence intensity (RFI). The RFI was generated as the ratio of the uorescence intensity between peri-pHin and peri-HdeA-72-DMN (with benzoate) at each pH point. (d) Calibration curve of membrane potential (DC) as a function of relative uorescence intensity (RFI) at pH 3. The RFI was generated as the ratio of the uorescence intensity between the red and green channels.

acid environment of the human stomach. The subsequent ow cytometric analysis and calculation showed cytosolic pH values of4.5 and 4.4, respectively, which were similar to the pH value whenE. coli was surrounded by pH 3 citrate buffer (Supplementary Fig. 20). Taken together, these results indicated that the digestive enzymes and inorganic salts present in biological systems were not interfering with our pH indicators.

PMF determination across the E. coli cytoplasmic membrane. To obtain the PMF value across the E. coli cytoplasmic membrane, we next employed a ratiometric uorescent probe for measurement of DC in E. coli cells by using ow cytometry.

DiOC2(3) is a lipophilic cyanine dye with a delocalized positive charge38,39. Upon excitation by a 488-nm laser, the uorescence of the stained cells could be detected in both the green channel (530 nm) and red channel (4600 nm). The green uorescence is dependent on cell size but independent of DC; while the red uorescence, due to the formation of dye aggregates, is dependent on both cell size and DC. The ratio of red to green uorescence would thus provide an accurate measurement of membrane potential. To calibrate the values of the calculated uorescence ratio to that of DC, the DiOC2(3)-stained E. coli cells were treated with potassium-specic ionophore, valinomycin, in the presence of different concentrations of external potassium ions38,40. Membrane potential was calculated from K distribution across the cytoplasmic membrane by means of the Nernst equation41. In this way, we generated the calibration curves of relative uorescence intensity (RFI, red uorescence intensity/ green uorescence intensity) as a function of DC at the extracellular pH 7 and pH 3 (Supplementary Fig. 21 and Fig. 4d). At pH 7, the RFI was a valid indicator of DC in the

range of 105 mV through 145 mV; whereas the RFI DC

was accurate between 40 and 80 mV at pH 3. On the basis of

the ow cytometric analysis of DiOC2(3)-stained E. coli cells both under neutral and acidic extracellular conditions, we determined that the membrane potential of E. coli was 124 mV when the

environment was at pH 7, which is consistent with previous reports41. In contrast, DC decreased to 47 mV when the

extracellular acidity dropped to pH 3, probably due to the acid-stress-induced depolarization of E. coli cell membrane42.

Finally, we calculated the PMF across E. coli cytoplasmic membrane under both neutral and acidic conditions. According to the equation PMF(mV) DC 59 DpH in Table 2, the PMF

was calculated as 159 mV at pH 7 (Supplementary Fig. 21). In

contrast, when the extracellular acidity dropped to pH 3, the PMF value was calculated to be 112 mV, which is smaller in

magnitude than that at pH 7. Therefore, when the external pH decreased from neutral to pH 3, the DC across the E. coli cytoplasmic membrane dramatically decreased from 124 to 47 mV, whereas the DpH values increased about 1.1 pH unit. Together, the PMF value across the E. coli cytoplasmic membrane underwent a relatively small decrease from 159 to 112 mV,

which is still highly negative.

DiscussionUsing a newly developed Cu(I) ligand in assisting the CuAAC reaction, we have achieved highly efcient protein labelling inside the bacterial cytoplasm without apparent toxicity. The tris(triazolylmethyl)amine-based ligand BTTP signicantly increased the redox potential of the coordinated Cu(I), sequestered Cu(I)-associated toxicity and dramatically accelerated the Cu(I)-catalysed 1,3-dipolar cycloaddition between azide- and alkyne-

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Table 2 | PMF of Escherichia coli cells under extreme acidic conditions (pH 3).

RFIcyto RFI pH DpH DW(mV) PMF(mV)

peri-pH 1.86 3.3 1.1 47 112

cyto-pH 0.98 1.04 4.4 RFI FI

peri-pHin/FIperi-HdeA-72-DMN (i)

RFIcyto FI

cyto-pHin/FIcyto-HdeA-72-DMN; RFI RFIcyto/0.94 (ii)

PMF, proton motive force; RFI, relative uorescence intensity.

In the absence of benzoate treatment, when the environmental pH is 3.0, the peri-RFI is calculated as the uorescence intensity (FI) ratio between peri-pHin and peri-HdeA-72-DMN (formula-i); the result of peri-RFI (1.86) corresponds to pH 3.3. Cyto-RFI is calculated as the FI ratio between cyto-pHin and cyto-HdeA-72-DMN and then converted to the calibration RFI (formula-ii); the result (1.04) corresponds to pH 4.4. DpH pH pH .

tagged molecules. By applying the power of this chemical transformation for compartment-specic protein labelling with a solvatochromic uorophore, we developed a new technique for measuring acidity in different intracellular spaces of E. coli cells. When the extracellular pH dropped to 3, our cytoplasm- and periplasm-specic pH measurement showed an B1.1 pH unit transmembrane pH gradient (DpH), which, in conjunction with the calculated membrane potential, allowed us to obtain the PMF value across the E. coli cytoplasmic membrane under this highly acidic condition. Our study revealed that E. coli cells actively maintain a pH gradient as well as a highly negative PMF value across its cytoplasmic membrane even under highly acidic conditions, which may be essential for acid-tolerance in Gram-negative bacteria. In addition, since PMF has important roles in energy production, cellular metabolism as well as bacterial motility, maintaining a relatively stable PMF in E. coli membrane under normal and stress conditions could be crucial in supporting diverse bacterial functions.

We have recently extended the genetic code expansion strategy into a panel of pathogenic enteric bacteria species including Shigella and Salmonella43. These two pathogens developed effective but different acid-resistance systems to survive through the highly acidic mammalian stomach to cause infections in the small intestine. Transferring our pH indicator system into these pathogenic species for compartment-specic pH measurement under acid stress may shed light on their acid-resistance mechanisms.

Methods

General materials. Bacterial cells were grown in LB (Luria-Bertani) medium. Antibiotics were used at nal concentrations of 50 mg ml 1 for ampicillin and35 mg ml 1 for chloramphenicol (Sigma). E. coli bacteria strain DH10B was used for the expression of GFP and HdeA variant proteins. Chemical compounds BPS,

TBTA, L-Histidine were purchased from Sigma-Aldrich, other compounds including ACPK, alk-4-DMN, BTTAA, BTTP and BTTPS were synthesized according to previous reports11,12,24,25.

Expression of GFP protein containing ACPK in E. coli. The plasmid pSupARMb-ACPK-RS was co-transformed with a plasmid carrying the GFP-149TAG gene (pBAD-GFP-149TAG) into E. coli DH10B cells. Bacteria were grown at 37 C in LB medium containing ampicillin (50 mg ml 1) and chloramphenicol (35 mg ml 1)

for 3 h till OD600 0.6, at which point 1 mM ACPK (nal concentration) was

added to the culture. The bacteria were continuously grown at 37 C for 30 min before being transferred to 30 C for induction in the presence of 0.2% arabinose for 10 h. Cells were collected by centrifugation (10 min, 6,000 r.p.m., 4 C). Proteins were extracted by sonication, and the extract was claried by centrifugation(30 min, 13,000 r.p.m., 4 C). The GFP protein containing ACPK was puried by HisTrap HP column (GE) operated with FPLC system (GE) according to standard protocols.

In vitro labelling of GFP-ACPK mutant and SDSPAGE analysis. GFP protein containing ACPK was buffered to phosphate-buffered saline (PBS) buffer (pH 7.4) and diluted to a protein concentration of 30 mM. A total of 500 ml of GFP-ACPK in

PBS buffer was reacted with alk-4-DMN (250 mM) at room temperature for 15 min in the presence of Cu(I)/ligand ([Cu] 50 mM, [ligand] 100 mM) and sodium

ascorbate (2 mM). The reaction mixture was diluted to 5 ml with PBS and concentrated to 500 ml using an Amicon Ultra 3,000 MWCO centrifuge lter (Sigma),

repeated four times to ensure that all the unreacted dye and Cu(I)/ligand were removed. The samples were analysed by SDSPAGE: a 415% gel was run at 160 V and was imaged using ChemDoc for ultraviolet absorption before Coomassie blue staining.

Biocompatible click-labelling inside bacterial cells. E. coli cells expressing GFP (or HdeA) protein variants carrying the site-specically incorporated ACPK handle were spun down and washed with PBS (pH 7.4) before being diluted into 0.5 ml PBS (OD600 0.6, containing 2% dimethyl sulphoxide). For the biocompatible

CuAAC reaction, Cu(I)ligand was added at a nal concentration of 100 mM (100 mM CuSO4 200 mM ligands), while the sodium ascorbate was added at a

nal concentration of 2.5 mM. The nal concentration of alk-4-DMN was 300 mM. The reaction was allowed to proceed at room temperature for 1 h with agitation and then quenched with bathocuproine disulphonate (5 mM). Cells were washed with PBS several times, and the samples were lysed for SDSPAGE analysis or diluted in different buffers for ow cytometry analysis.

In vitro detection of ROS production using coumarin-3-carboxylic acid. A total of 5 mM coumarin-3-carboxylic acid was prepared as stock solution in DMF (N,N-dimethylformamide). CuSO4 or CuSO4-ligands were prepared in PBS buffer at a concentration of 100 mM (200 mM ligands), then 2 mM sodium ascorbate was added. The mixture was incubated at room temperature for 15 min, then cou-marin-3-carboxylic acid was added into the reaction buffer (100 mM). The samples were excited at 395 nm and the emission spectra were recorded from 400600 nm. Cu(I)BPS complex showed uorescence upon 395-nm excitation, therefore it was not used in this assay.

In vivo detection of ROS production using DCFH-DA. The Cu(I)ligands-treated (100 mM CuSO4 200 mM ligands, 2 mM sodium ascorbate, incubated at room

temperature for 1 h) E. coli cells were collected and resuspended in PBS buffer (OD600 0.6), 5 mM (nal concentration) of DCFH-DA (1 mM stock in dimethyl

sulphoxide, Applygen Technologies Inc) were added to the cultures and incubated at room temperature in the dark for 30 min. DCFH-DA is a nonpolar dye which is converted into the polar derivative DCFH by cellular esterases. DCFH is nonuorescent but switched to highly uorescent DCF when oxidized by intracellular ROS or other peroxides, DCF has an excitation wavelength of 485 nm and an emission band between 500 and 600 nm. After incubation samples were transferred into a 96-well round-bottomed culture plate, and the uorescence intensity was measured using a Synergy Hybrid plate reader (excitation: 48515 nm, emission: 53015 nm).

Fluorescent measurements of Cu(I)ligand-treated GFP. GFP protein containing ACPK were buffered in PBS and diluted into a concentration of 30 mM (50 ml). Free Cu(I) or Cu(I)ligand complex (50 mM CuSO4, 100 mM ligand) and 2 mM sodium ascorbate were added, and incubated at room temperature for30 min. The Cu(I) was then removed by Micro Bio Spin-6 desalting column (Bio-Rad). The proteins were then diluted to 500 ml PBS buffer, transferred into a 96-well round-bottomed culture plate, and the uorescence intensity was measured using a Synergy Hybrid plate reader (excitation: 48015 nm, emission:52015 nm).

PI staining assay of Cu(I)-treated E. coli cells. The Cu(I)ligands-treated (100 mM CuSO4 200 mM ligand, 2 mM sodium ascorbate, incubation at room

temperature for 1 h) E. coli cells were collected and resuspended in PBS buffer (OD600 0.6), stained with PI (3 mg ml 1) and incubated at room temperature for

15 min. Cells were then washed with PBS three times, transferred into a 96-well round-bottomed culture plate and the uorescence intensity was measured using a Synergy Hybrid plate reader (excitation: 53520 nm, emission: 61520 nm).

Flow cytometry. Analysis of cells by ow cytometry was carried out on a BD FACSCalibur Flow Cytometer equipped with a 488-nm laser. Fluorescence was

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detected in FL-1 channel for alk-4-DMN and FL-3 channel for trypan blue, uorescence of DiOC2(3) was detected both in the FL-1 channel and FL-3 channel.

After uorescent labelling, the cells were washed with PBS four times and then diluted 1:1,000 in their respective buffers for uorescence determination. Between 5 104 and 1 105 events were collected for each experiment; three parallel

experiments were carried out for each sample. Data analysis was performed with CellQuest Pro software (BD Biosciences).

Isolation of periplasmic and cytosolic proteins. E. coli cells (in 10 ml culture) were collected by centrifugation at 6,000 r.p.m. for 5 min at 4 C (ref. 44). The pellets were then resuspended in 0.5 ml buffer containing 10 mM Tris (pH 7.4),1 mM EDTA, 15,000 U ml 1 Lysozyme, 20% (w/v) sucrose and incubated at room temperature for 10 min, 0.5 ml ice-cold water was then added and the sample was incubated on ice for an additional 5 min. Spheroplasts were collected by centrifugation (12,000 r.p.m., 5 min, 4 C), and the supernatant was collected as the periplasmic fraction (peri-). The spheroplasts were resuspended in 0.5 ml buffer containing 10 mM Tris (pH 7.4), 1 mM EDTA and sonicated on ice for 5 min, and after centrifugation (12,000 r.p.m., 10 min, 4 C), the supernatant was collected as the cytoplasmic fraction (cyto-). The samples were analysed by SDSPAGE. Rabbit anti-HdeA (raised from rabbits)45 and mouse anti-EF Tu (Santa Cruz Biotechnology) were used to verify this method.

Electrochemical measurement. Cu(I) complexes in ddH2O were made in situ by directly mixing the proper amounts (70 ml, 500 mM nal concentration) of Cu(CH3CN)4PF6(I) solution (50 mM stock) in acetonitrile and the corresponding ligand in the cell (7 ml total volume) in a glove box under nitrogen35,46.

Voltammetric measurements were carried out with CHI-660D electrochemical work station (ShangHai ChenHua). Experiments were performed at room temperature using a saturated calomel electrode as the reference electrode. The working electrode was a Pt electrode (ShangHai ChenHua, 3 mM diameter) and polished before experiment. Cyclic voltammetry was performed in the potential range of 0.6 to 0.6 V versus SCE at 50 mV s 1.

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Acknowledgements

We are grateful to Professor Erik L. Snapp and Miss Samantha Wilner for valuable discussions. This work was partially supported by research grants from the National Key Basic Research Foundation of China (2010CB912302 and 2012CB917301) and National Natural Science Foundation of China (21225206 and 91313301) to P.R.C., the National Institutes of Health to P.W. (R01GM093282), and the National High Technology Research and Development Program of China (2014AA020512) to J.Z. Graduate fellowship funding to A.S.J. was provided by the NIH Training Program in Cellular and Molecular Biology and Genetics (T32 GM007491).

Author contributions

M.Y., A.S.J. and W.W. performed all the experiments. M.Y. and A.S.J. prepared the gures. P.R.C., P.W. and J.Z. conceived the study, analysed the data and wrote the manuscript, with edits from all the authors.

Additional information

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Competing nancial interests: The authors declare no competing nancial interests.

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How to cite this article: Yang, M. et al. Biocompatible click chemistry enabled compartment-specic pH measurement inside E. coli. Nat. Commun. 5:4981 doi: 10.1038/ncomms5981 (2014).

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