Live-cell labelling techniques to visualize proteins with minimal disturbance are important; however, the currently available methods are limited in their labelling efficiency, specificity and cell permeability. We describe high-throughput protein labelling facilitated by minimalistic probes delivered to mammalian cells by microfluidic cell squeezing. High-affinity and target-specific tracing of proteins in various subcellular compartments is demonstrated, culminating in photoinduced labelling within live cells. Both the fine-tuned delivery of subnanomolar concentrations and the minimal size of the probe allow for live-cell super-resolution imaging with very low background and nanometre precision. This method is fast in probe delivery (∼1,000,000 cells per second), versatile across cell types and can be readily transferred to a multitude of proteins. Moreover, the technique succeeds in combination with well-established methods to gain multiplexed labelling and has demonstrated potential to precisely trace target proteins, in live mammalian cells, by super-resolution microscopy.
Direct observation of intracellular processes has the potential to yield insight into fundamental biological pathways and disease mechanisms. Several techniques have been developed to enable high-resolution imaging of live cells; yet, the limited ability to trace intracellular components has hindered progress. Hence, two of the persistent challenges are probe design and cellular delivery with minimal toxicity, pivotal for advances in live-cell imaging technologies. Here we describe an efficient approach to tag and image intracellular components in live mammalian cells. Using the microfluidic cell squeezing platform to deliver small fluorescent tris-N-nitrilotriacetic acid (trisNTA) probes (∼1 kDa), we demonstrate highly efficient, minimally disruptive, light-triggered tracing of native proteins and the subsequent super-resolution imaging of live-cell phenomena.
Live-cell microscopy contributed significant knowledge of dynamic processes such as protein trafficking and single-molecule localization-based imaging techniques visualize proteins with high-resolution information (≤50 nm)1,2. All fluorescent imaging techniques require protocols to introduce the label with the need to minimize its influence on the system. Fluorescent proteins, self-labelling tags3,4,5,6 or labelling by enzymatic methods7,8 can interfere with protein function, assembly or dynamics. Bulky fusion proteins (>20 kDa) entail the risk of steric hindrance and functional perturbations, whereas smaller tags (for example, tetracysteine tag) deal with unspecific interactions or require additional experimental steps8 and optimized flanking sequences for each protein target9. Although synthetic fluorophores have enhanced photostability, quantum yield, spectral range and localization precision, it is difficult to introduce such probes to the cytosolic environment using existing delivery technologies. On the one hand, current transduction strategies such as delivery by cell-penetrating peptides (CPPs), electroporation and so on are suboptimal, suffering from poor and endosomal uptake, rapid degradation by extracellular and endosomal proteases, low in vivo efficiency or elaborated chemical synthesis. On the other hand, antibody-based labelling approaches, for example, are limited to chemically arrested (fixed) cells and the availability of specific antibodies for a protein target. Owing to the described limitations of existing labelling and transduction technologies, there is a persistent demand for techniques enabling high-throughput in-cell labelling by minimal tags that are conductive to high-resolution and super-resolution microscopy.
Here we demonstrate robust in-cell targeting of native proteins using a labelled multivalent chelator head trisNTA10 and a genetically encoded oligohistidine sequence (Fig. 1a). trisNTA site specifically recognizes His6–10-tagged proteins in the (sub)nanomolar range (Kd of 0.1–10 nM) even in the crowded cellular environment11. The minimal size of the tag and the molecular probe allows direct targeting with nanometre precision at subnanomolar concentrations as required for single-molecule localization-based imaging techniques1,2,12,13 with no impact on intracellular trafficking or demand for additional cofactors affecting endogenous processes. We simplified efficient transfer of the trisNTA probe into living cells by cell squeezing14, combining precisely controlled cytosolic delivery with high specificity and low cytotoxicity. Briefly, transient cell permeabilization is achieved by rapid viscoelastic deformation of cells as they pass through micrometre-scale constrictions. This facilitates fast uptake of probes into the cytosol before cell-intrinsic repair mechanisms kick in15.
High-affinity protein labelling at subnanomolar concentration
We first investigated the specificity of the trisNTA/His tag targeting in chemically arrested cells. To evaluate precise localization, different proteins resident at distinct subcellular compartments were selected: (i) the transporter associated with antigen processing (TAP) in the membrane of the endoplasmic reticulum16; (ii) histone 2B (H2B) in the nucleus; and (iii) Lamin A at the nuclear envelope. All proteins of interest (POIs) were fused to a His10 tag and a fluorescent protein (TAP1mVenus-His10, H2BmVenus-His10 and His10-mEGFPLamin A) for specific targeting and co-localization studies, respectively. For sensitive detection, trisNTA was covalently coupled to different fluorescent dyes (trisNTAf, f=Alexa488, ATTO565, ATTO647N, Alexa647 and ATTO655). Mammalian cells were transiently transfected with the corresponding target genes. His-tagged proteins were specifically stained by trisNTAf with excellent co-localization and signal-to-noise ratio (Pearson’s coefficients between 0.90 and 0.96), using confocal laser scanning microscopy (CLSM; Fig. 1b and Supplementary Fig. 1). Strikingly, even at 200 pM of trisNTAf, His-tagged proteins were labelled with high specificity (Supplementary Fig. 2). By analysing a variety of fluorescent dyes, we noticed that trisNTAATTO565 targeting produced a higher background compared with trisNTAATTO647N, trisNTAAlexa647 or trisNTAATTO655 (Supplementary Fig. 3). This was assigned to unspecific binding of the ATTO565 dye. Moreover, the superposition of both fluorescence intensity profiles reflects an excellent correlation between the POI expression level and the labelling density of trisNTAAlexa647 (Fig. 1c,d, Pearson’s coefficient r=0.95). Notably, using nanomolar concentrations, trisNTAf labelling is significantly more efficient within 30 min than SNAPf-tag labelling (Supplementary Fig. 4). In contrast, mammalian cells expressing H2B lacking a His tag showed neither trisNTAf labelling nor unspecific staining (Supplementary Fig. 5). In conclusion, trisNTAf targeting at subnanomolar concentrations is highly specific to trace His-tagged proteins. To exploit these benefits further, we combined trisNTAf with well-established labelling methods for multiplexed protein modification. Specific trisNTAAlexa488 labelling of His10LaminA (Fig. 1e, green) was successfully achieved in combination with SNAPf-tag labelling of H2B (magenta) and antibody labelling of tubulin (red), as well as the lysosomal-associated membrane protein 1 (blue). Thus, the ultra-small interaction pair complements the toolbox of well-established labelling techniques and the nanomolar concentrations perform various avenues in multiplexed labelling.
High-throughput live-cell labelling within mammalian cells
Encouraged by these observations, we aimed at protein labelling in living cells. To transfer trisNTAf into cells, we applied microfluidic cell squeezing (Fig. 2a). As the trisNTAf probes are chemically diverse relating to the used fluorophores, common transduction strategies are unlikely to efficiently deliver nanomolar concentrations of trisNTAf into mammalian cells. Specifically, mammalian cells were mechanically pushed (‘squeezed’) through micrometre constrictions at elevated pressure of 30 psi. This approach allows for high cell survival (>90%) and efficient uptake of trisNTAf (up to 80%; Supplementary Figs 6 and 7). Energy-dependent endocytosis, often observed at cargo transfer with supercharged molecules (Supplementary Fig. 8) or low concentrations of CPPs (Supplementary Fig. 9)17, were prevented by performing cell squeezing at 4 °C (Supplementary Fig. 10).
By squeezing TAP1mVenus-His10-transfected HeLa cells in the presence of trisNTAf (100 nM), we achieved a high-throughput delivery and a high-density labelling, illustrated by an excellent co-localization between both reporter molecules (Fig. 2b). We noticed that both probe delivery by cell squeezing and protein labelling are highly reproducible (n>20). To quantify the trisNTAf delivery, we performed flow cytometry analysis on micromanipulated cells. Thirty-five per cent of transfected cells (54% of total) were effectively transduced with trisNTAATTO565 (Fig. 2c), which is > 30 × more efficient for trisNTAf delivery than electroporation (Supplementary Fig. 11). Squeezing of up to 1,000,000 cells per second enables live-cell labelling at high throughput and reproducibility, and hence largely exceeds the efficiency achieved by other direct transfer methods such as microinjection. The massively parallel and constant cell transduction surpasses the stochastic, while variable efficient uptake by CPPs (Supplementary Fig. 9) and is more than 1,000-fold below the micromolar probe concentrations used by elegant self-labelling tags, for example, SNAP tag3,4,5,6.
Highly specific protein targeting with minimal disturbance
Analogous micromanipulation and live-cell labelling was applied to H2BmVenus-His10- and His10-mEGFPLamin A-transfected cells (Fig. 2d). Cell transduction was analysed 15 min and 1 h after squeezing by CLSM. An excellent co-localization between the trisNTAf reporter molecule and the His-tagged POIs was observed (Pearson’s coefficients range from 0.81 to 0.93), in line with the subcellular localization in chemically arrested cells. Notably, live-cell labelling is independent of cell types, for example, HeLa, HeLa Kyoto, Chinese hamster ovary (CHO-K1) or human embryonic kidney 293 cells (Figs 1b and 2d). Beyond that, trisNTAf labelling after removal of the bulky fluorescent protein fully exploited the small size of the lock-and-key element and confirmed again specific labelling in living cells with minimal perturbation (Supplementary Fig. 12). The specificity was validated in cells transfected with H2BEGFP lacking a His tag and untransfected cells (Supplementary Fig. 13). In both cases, neither co-localization of trisNTAf with H2BEGFP nor unspecific binding was detected. Delivery of nickel-free trisNTAf or of free Alexa647 dye showed no labelling in TAP1mVenus-His10-transfected cells (Supplementary Figs 14 and 15). In contrast, trisNTAAlexa647 (Fig. 2 and Supplementary Fig. 14) and trisNTAATTO565 (Supplementary Fig. 16) clearly stain TAP1mVenus-His10 at the endoplasmic reticulum membrane after cell squeezing. Noticeably, cell viability of trisNTAf-transduced cells was negligibly affected 1 and 24 h after labelling. Similar concentrations of unbound nickel ions inside mammalian cells had no significant toxic effects (Supplementary Fig. 7). In contrast, electroporation entailed more than twofold increased toxicity compared with squeezing (Supplementary Figs 7 and 11). Collectively, nanomolar delivery of trisNTAf fully realized the potential of in-cell protein manipulation with minimal perturbation and modification rates exceeding common approaches.
After successful in-cell labelling of different His-tagged proteins, we aimed for in vivo multiplexed labelling by combining trisNTAf with well-established labelling methods. By trisNTAAlexa647 delivery via squeezing and subsequent SNAPf-tag labelling, we achieved specific and distinct targeting of His10LaminA in the presence of two different SNAPf-tagged proteins in live cells (Fig. 3a and Supplementary Fig. 17). Hence, trisNTAf complements the toolbox for in vivo multiplexed labelling, offering minimal disturbance due to its small size and simultaneously using low nanomolar concentrations.
We next determined the minimal reporter concentration required for specific live-cell labelling. Well-resolved images of TAP1mVenus-His10 were obtained even at 1 nM of trisNTAAlexa647 (Fig. 3b). Based on previous observations, approximately one-third of the cargo provided during squeezing is the effective intracellular concentration14. Thus, the estimated cytosolic concentration of trisNTAf further corroborates the high target sensitivity at subnanomolar concentrations (∼300 pM). These results are in line with the detection limit of ∼200 pM trisNTAAlexa647 in chemically arrested cells (Supplementary Fig. 2). Hence, this enables the precise adjustment of the effective, intracellular trisNTAf concentration to improve the signal-to-background ratio, hardly realized by alternative approaches at nanomolar probe concentrations (for example, CPPs, SNAP, CLIP and Halo tag; Supplementary Figs 4 and 9)3,4,6,18, and circumvents endocytic uptake observed with supercharged proteins at similar nanomolar concentrations (Supplementary Fig. 8)19.
In-cell protein modification with nanometre precision
Incited by this observation, we aimed at temporal and spatial control of protein tracing by light, which depends on low probe concentrations for high signal-to-background ratios. Using photoactivatable trisNTAf (PA-trisNTAATTO565)20 for dynamic cellular imaging on demand, light-activated in vivo labelling of His10-mEGFPLamin A was demonstrated up to 24 h after squeezing (Fig. 3c). Notably, already a 10-s 405-nm light pulse sufficiently activated PA-trisNTA at single-cell level and led to excellent co-localization in a dose-adapted manner. This probe enables in situ labelling at defined time points such as certain mitotic phases and paves the way for live-cell protein tracing with high temporal resolution. The nanomolar concentrations (≤10 nM) and in particular the small size of the tag and probe are especially beneficial for advanced microscopy techniques, bringing the fluorophore in 1-nm proximity to the target protein. Hence, we performed live-cell super-resolution microscopy with trisNTAATTO655 on His10-mEGFPLamin A-transfected cells. Using direct stochastic optical reconstruction microscopy (dSTORM)21, Lamin A structures with a high signal-to-noise ratio were obtained in the super-resolved images of live cells. A localization precision of 16.4±3.1 nm was achieved, resulting in a resolution of 40 nm by in-cell trisNTAATTO655 tracing with substantially increased resolution compared with diffraction-limited fluorescence microscopy (Fig. 3d).
We established a high-throughput method for protein labelling inside living cells using a minimalistic lock-and-key probe. Our method is versatile in the choice of the molecular probe, cell type and the subcellular localization of the POIs, a persistent challenge in live-cell analysis. The high-affinity trisNTA/His tag interaction pair enables fast labelling (≤10 min) at subnanomolar concentrations with tunable labelling density and flexibility of cell-impermeable organic fluorophores. Compared with carrier-mediated transport by CPPs11,18, delivery of 1,000-fold lower concentrations (nM versus μM) effectively decreases the fluorescence background. Furthermore, high-throughput analysis with up to 1,000,000 cells per second can be achieved in contrast to microinjection. In addition, trisNTAf delivery via squeezing avoids endocytic cargo uptake, frequently observed with low CPP concentrations and supercharged molecules, offering decreased toxicity and a >30-fold higher efficiency compared with electroporation. The minimal probe complements the toolbox of well-established labelling techniques such as self-labelling enzymes and can be combined with the latter to achieve distinct labelling of different proteins in fixed as well as in living cells. Moreover, in situ photoactivation of PA-trisNTA allows labelling at defined time points, to trace proteins for dynamic cellular imaging. The achieved close target proximity of the labelling pair substantially improved the localization accuracy in live-cell super-resolution microscopy. Remarkable aspects of our approach are the speed, flexibility and efficiency for high-throughput live-cell targeting of proteins even if assembled in stable and transient macromolecular complexes. This study is exploited via one of the smallest high-affinity lock-and-key recognition pairs known so far and allows even multiple cargos to be delivered simultaneously, displaying diverse chemical properties. The quantity of cargo for in-cell manipulation can be precisely tuned and the biological output can in turn be fine-tuned. As the affinity tag is widely used in life sciences and our delivery platform is broadly applicable across cell types, this live-cell labelling method could potentially be implemented across numerous cell-impermeable probes and prodrugs, as well as translated to difficult cell lines including patient-derived cells and embryonic stem cells, providing the opportunity to use these cells for advanced microscopy techniques and live-cell analysis.
The H2B construct H2BmVenus-His10 was generated by consecutive insertion of H2B and mVenus-His10 into pCDNA3.1(+) (Life Technologies). mVenus-His10 was PCR amplified using Phusion High-Fidelity DNA Polymerase (Fermentas) and the primer pair forward (fw) 5′-GCGCGCGCGGCCGCGTGAGCAAGGGCGAGGAGCTGTTCA-3′ (NotI restriction site underlined) and reverse (rev) 5′-GCGCGCTCTAGATTAGTGATGGTGGTGATGATGATG-3′ (XbaI restriction site underlined), and cloned into the pCDNA3.1(+) plasmid using the indicated restriction enzymes (Fermentas). Subsequently, H2B was amplified using the primer pair fw 5′-GCGCGCGGTACC ATGCCAGAGCCAGCGAAGTCTGCTCCCGC-3′ (Acc65I restriction site underlined, start codon bold) and rev 5′-GCGCGCGCGGCCGCTCTTGG AGCTGGTGTACTTAGTGAC-3′ (NotI restriction site underlined), and cloned into the pCDNA3.1(+) plasmid, amino terminally of mVenus-His10. As a template for the amplification of H2B, the plasmid pEGFP-N1 containing the human H2B sequence (plasmid 11680, Addgene) was used, which also served as control for the specificity of trisNTAf for His-tagged POIs in living cells (Supplementary Figs 5 and 13). The plasmid encoding for human Lamin A was generously provided by Dr Sascha Neumann (Institute of Biochemistry, University of Cologne)22 and used as template to amplify Lamin A with the primer pair fw 5′-GCGCGCCTCGAGCTATGGAGACCCCGTCCCAGCGGCGCGCCACCCG-3′ (XhoI restriction site underlined) and rev 5′-GCGCGCGATATC TTACATGATGCTGCAGTTCTGGGGGCTCTGGG-3′ (EcoRV restriction site underlined, stop codon bold). Lamin A was inserted into the pcDXC3GMS plasmid (Addgene) already containing His10-mEGFP, leading to the fusion gene encoding for His10–mEGFPLamin A. To generate the plasmid containing His10Lamin A without a fluorescent protein, Lamin A was amplified via PCR, simultaneously introducing a His10-tag with the primer pair fw 5′-GCGCGCGGATCCACCATG CACCATCATCATCATCATCACCACCATCACTCCGGACTCAGATCTCGAGTCATGGAGACCC-3′ (BamHI restriction site underlined, start codon bold, His10-tag italic) and rev 5′-GCGCGCGCGGCCGC TTACATGATGCTGCAGTTCTGGGGGCTCTGGG-3′ (NotI restriction site underlined, stop codon bold). Indicated restriction sites were used to insert this PCR product into the pCDNA3.1 (+) vector. The pSNAPf-H2B and pSNAPf-Cox8A plasmids (New England Biolabs) were used for SNAPf-tag labelling. In addition, a plasmid coding for the core domain of TAP1, tagged with mVenus-His10 (TAP1mVenus-His10) was used as previously described23.
Cell culture and transfection
HeLa cells, HeLa Kyoto cells, Chinese hamster ovary (CHO-K1) cells and human embryonic kidney 293 cells were maintained in DMEM medium with 4.5 g l−1 glucose (Gibco), supplied with 10% (v/v) FCS (Gibco) in T75 cell culture flasks (Greiner). Every 2–3 days, cells were passaged using PBS (Sigma-Aldrich) and 0.5% trypsin/0.02% EDTA/PBS (GE Healthcare). All cell lines were cultivated in a humidified tissue culture incubator at 37 °C and 5% CO2. Mycoplasma contamination tests were carried out regularly, following the guidelines described24. Transient transfection was performed with Lipofectamine 2000 (Life Technologies), following the manufacturer’s instructions. For fixation and staining, 2 × 104 cells per well were seeded into eight wells on cover glass II slides (Sarstedt) and transfected with 0.2 μg DNA per well. For squeezing experiments, 8 × 105 cells were seeded into six-well cell culture plates (Greiner) and transfected with 2 μg DNA per well. After transfection, cells were incubated 12–48 h at 37 °C and 5% CO2 until experiments were performed.
Cell viability test
To analyse cell viability after squeezing, the Sytox Blue Dead Cell stain (Life Technologies) was used to stain cells with a permeable plasma membrane. Cells were squeezed in the presence of 100 nM trisNTAAlexa647, 100 nM Alexa647 dye or 500 nM NiCl2, followed by incubation with 1 μM of Sytox Blue Dead Cell Stain (20 min, room temperature (RT)) at different time points. Cell viability analysis was performed using the Attune flow cytometer (Life Technologies) and data were processed using FlowJo 7.6.5 (Tree Star Inc.). Before detaching the cells, the supernatant was collected to avoid altering the results by removing dead cells during washing. Identical Sytox Blue Dead Cell Staining was conducted with cells after electroporation (described above), followed by flow cytometry analysis of cell viability and uptake of trisNTAATTO565. All experiments were performed in triplicates and error bars indicate the s.d.
Imaging was performed using the confocal laser scanning TCS SP5 microscope (Leica) and a Plan-Appochromat 63 × 1.4 Oil differential interference contrast objective. Images were acquired sequentially to avoid cross-talk. The following laser lines were used for excitation: 405 nm (diode laser) for 4,6-diamidino-2-phenylindole, Hoechst and Sytox Blue Dead Cell Stain; 488 nm (argon laser) for monomeric enhanced green fluorescent protein (mEGFP), Alexa488, MitoTrackerGreen and Syto16; 488 or 514 nm (argon laser) for mVenus; 565 nm (diode-pumped solid-state laser) for ATTO565 and TMR-Star; and 633 nm (helium-neon laser) for ATTO647N, Alexa647 and ATTO655. For image analysis, ImageJ25 was used in combination with Fiji26.
A custom-built microscope was used for super-resolution imaging of trisNTAATTO655-labelled His10-mEGFPLamin A27. Samples were illuminated with 488 nm (Sapphire 488 LP, Coherent) and 643 nm (iBeam smart, Toptica Photonics) laser beams in total internal reflection fluorescence mode. The excitation light was focused on the back focal plane of a 100 × oil objective (PLAPO 100 × , total internal reflection fluorescence mode, numerical aperture 1.45, Olympus) mounted on an inverted microscope (Olympus IX71). The emission was recorded using an electron multiplying charge-coupled device camera (Ixon3, Andor) with frame-transfer mode, 5.1 × pre-amplifier gain and electron multiplying (EM) gain set to 200. For every sample, 40,000 images were recorded at a frame rate of 33 Hz and image reconstruction was performed with rapidSTORM28. The localization (σloc) precision of the dSTORM images was calculated to 16.4±3.1 nm and a resolution of ≤40 nm (for dSTORM image see Fig. 3d). Calculations were performed according to Mortensen et al.29
Fixation and trisNTAf labelling
Before fixation, cells were washed with PBS (Sigma-Aldrich). Fixation with 4% formaldehyde (Roth)/PBS for 15 min at RT was followed by quenching using 50 mM glycine/PBS (10 min, RT; Roth) and permeabilization with 0.1% Triton X-100/PBS (10 min, RT; Roth). After blocking with 5% (w/v) BSA (Albumin Fraction V, Roth) in PBS (1 h, RT), cells were stained with 100 nM of trisNTAf in 1% (w/v) BSA/PBS. Cells were stained with 0.1 μg ml−1 4′,6-diamidino-2-phenylindole (Sigma-Aldrich) in 1% BSA/PBS for 30 min–1 h at RT. After washing with 5% BSA/PBS (3 × ), cells were postfixed with 2% formaldehyde/PBS (15 min, RT) and stored in PBS until confocal imaging was performed. When trisNTAf labelling was combined with antibody labelling, 100 nM trisNTA was applied together with the primary antibodies for 1 h at RT. After subsequent washing with PBS, cells were incubated with secondary antibodies. Antibodies were all diluted in 1% BSA/PBS as follows: rabbit α-tubulin (Abcam, 1:350), mouse α-human CD107a (LAMP-1; Biolegend, 1:1,000), goat α-rabbitCy3 (Abcam, 1:1,000) and goat α-mousePacificBlue (Life Technologies, 1:1,000). trisNTA was conjugated with the corresponding fluorescent dyes by reacting amino-trisNTA with the N-hydroxysuccinimide (NHS)-activated dye, respectively. After RP-C18 HPLC chromatography, the multivalent chelator head was loaded with Ni2+ (10 mM NiCl2 in 20 mM HEPES, pH 7.0) and purified by anion exchange chromatography (HiTrap Q HP; GE Healthcare)11.
SNAPf-tag labelling in fixed cells
HeLa Kyoto cells transfected with H2BSNAPf were labelled with O6-benzyl guanineAlexa647 (New England Biolabs) 24 h after transfection. After washing with PBS, cells were fixed with 4% formaldehyde/PBS (10 min, RT), washed twice with PBS and subsequently permeabilized using 0.1% Triton X-100/PBS (10 min, RT). Cells were washed three times with 0.5% BSA/PBS and incubated with 10 or 100 nM of benzyl guanineAlexa647 for 30 min at 37 °C. After three washing steps with 0.1% Triton X-100/0.5% BSA/PBS and two washing steps with PBS, imaging was conducted by CLSM. In case of combined trisNTAf- and SNAPf-tag labelling, SNAPf-tag labelling was performed first, followed by trisNTAf labelling as described above.
trisNTAf delivery via cell squeezing
Squeezing was performed using a chip with constrictions of 7 μm in diameter and 10 μm in length (CellSqueeze 10-(7)x1, SQZbiotech), if not otherwise stated. In all microfluidic experiments, a cell density of 1.5 × 106 cells per ml in 10% (v/v) FCS/PBS were squeezed through the chip at a pressure of 30 psi. Transduction was conducted at 4 °C, to block cargo uptake by endocytosis. During squeezing, the following cargo concentrations were used: 0.1–100 nM of trisNTAf (f=ATTO565, ATTO647N, Alexa647 or ATTO655), 200 nM of PA-trisNTAATTO565 and 100 nM of free Alexa647 (Molecular Probes). After squeezing, cells were incubated for 5 min at 4 °C, to reseal the plasma membrane15. Squeezed cells were washed with DMEM containing 10% FCS and 10 mM histidine (Sigma-Aldrich), to remove unspecifically bound trisNTAf from the cell surface, seeded into eight wells on cover glass II slides (Sarstedt) in DMEM containing 10% FCS and cultured at 37 °C and 5% CO2. Confocal imaging was performed at different time points (0.25, 0.5, 1, 2 and 24 h) after squeezing. As a control for endosomal uptake, cells were incubated with 100 nM of trisNTAf at RT and 4 °C without microfluidic cell manipulation (Supplementary Fig. 10). In case of combined trisNTAf and SNAPf-tag labelling, cells were first squeezed in the presence of 100 nM trisNTAAlexa647 and 3 μM SNAP-Cell TMR-Star (New England Biolabs) was added 5 min after squeezing. After 15 min incubation at 37 °C and 5% CO2, cells were washed with 10% FCS/DMEM and incubated again 30 min under standard cell culture conditions before CLSM imaging was performed. To visualize mitochondria in case of Cox8ASNAPf labelling MitoTrackerAlexa488 was added 10 min prior imaging.
trisNTAf delivery by the CPP Tat49–57
TAPmVenus-His10-transfected HeLa Kyoto cells were incubated with different concentrations (10 μM and 100, 10 and 1 nM) of a non-covalent complex composed of Tat49–57His6 and trisNTAAlexa647 for 30 min. After washing with 10% FCS/DMEM and PBS at 37 °C, trisNTA uptake and labelling of His10-tagged TAP was analysed by live-cell imaging via CLSM11.
trisNTAf delivery via supercharged GFP (GFP36+)
trisNTAATTO565 (100 nM) and His6GFP36+ (100 nM) were incubated for 30 min at RT to form the trisNTA-His tag complex. HeLa cells were washed with PBS and treated with the pre-formed complex at 37 °C. In vivo uptake was immediately followed by CLSM. After 20 min, cells were washed three times with PBS and 20 U ml−1 heparin/PBS (2 × ), to remove the complex from the plasma membrane. Internalization of trisNTA/His6GFP36+ was analysed by CLSM after washing three times with PBS. His6GFP36+ was expressed in BL21(DE3) Escherichia coli. After lysis by sonication in 2 M NaCl/PBS, His6GFP36+ proteins were purified via immobilized metal ion affinity chromatography using Ni Sepharose 6 Fast Flow (GE Healthcare). Elusion was performed with 500 mM imidazole before desalting of the eluted protein was conducted with PD-10 desalting columns (GE Healthcare)19.
trisNTAf delivery via electroporation
Trypsinized HeLa Kyoto cells were permeabilized via electroporation in the presence of trisNTAATTO565 (100 nM) with a Nucleofector Device (Lonza) using Nucleofector Kit V and program I-013. Transduced cells were washed with 10% FCS/DMEM and plated into eight-well cover glass II slides. To quantify trisNTAf uptake, flow cytometry analysis (as described above) and CLSM imaging was performed 1 h after electroporation. Before CLSM imaging, transduced HeLa cells were incubated with Sytox Blue Dead Cell Stain or Syto16 (Life Technologies) live cell stain, to distinguish between dead and live cells. Cell viability was analysed by flow cytometry using Sytox Blue Dead Cell Stain (see above).
Photoactivation of PA-trisNTAf
HeLa Kyoto cells, transfected with His10-mEGFP Lamin A, were squeezed in the presence of PA-trisNTAATTO565 (200 nM)20 and incubated at 37 °C and 5% CO2. Imaging via CLSM and photoactivation was conducted 2 or 24 h after squeezing, after cells reattached to the glass surface. PA-trisNTAATTO565 was photoactivated in single cells by illumination with the 405-nm laser for 10 s. Imaging of both channels (mEGFP and ATTO565) was performed before and directly after photoactivation.
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The German Research Foundation (Cluster of Excellence—Macromolecular Complexes to R.W., M.H. and R.T., as well as CRC 807, SPP 1623 and RTG 1986 to R.T. and SFB 807 to M.H.) supported the work. We thank Drs Sascha Neumann (Institute of Biochemistry, University of Cologne, Germany) and Ulrich Rothbauer (The Natural and Medical Sciences Institute, University of Tübingen, Germany) for generously providing us with the original Lamin A construct and the HeLa Kyoto cells, respectively. Furthermore, we thank Valentina Herbring and Dr Peter Mayerhofer for help with flow cytometry, and Markus Braner for helpful suggestions on the manuscript.
The authors declare no competing financial interests.
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Kollmannsperger, A., Sharei, A., Raulf, A. et al. Live-cell protein labelling with nanometre precision by cell squeezing. Nat Commun 7, 10372 (2016). https://doi.org/10.1038/ncomms10372
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