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. 2023 May 11;62(11):1735–1743. doi: 10.1021/acs.biochem.2c00712

Orthogonal Versatile Interacting Peptide Tags for Imaging Cellular Proteins

Alexa Suyama 1, Kaylyn L Devlin 1, Miguel Macias-Contreras 1, Julia K Doh 1, Ujwal Shinde 1, Kimberly E Beatty 1,*
PMCID: PMC10249344  PMID: 37167569

Abstract

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Genetic tags are transformative tools for investigating the function, localization, and interactions of cellular proteins. Most studies today are reliant on selective labeling of more than one protein to obtain comprehensive information on a protein’s behavior in situ. Some proteins can be analyzed by fusion to a protein tag, such as green fluorescent protein, HaloTag, or SNAP-Tag. Other proteins benefit from labeling via small peptide tags, such as the recently reported versatile interacting peptide (VIP) tags. VIP tags enable observations of protein localization and trafficking with bright fluorophores or nanoparticles. Here, we expand the VIP toolkit by presenting two new tags: TinyVIPER and PunyVIPER. These two tags were designed for use with MiniVIPER for labeling up to three distinct proteins at once in cells. Labeling is mediated by the formation of a high-affinity, biocompatible heterodimeric coiled coil. Each tag was validated by fluorescence microscopy, including observation of transferrin receptor 1 trafficking in live cells. We verified that labeling via each tag is highly specific for one- or two-color imaging. Last, the self-sorting tags were used for simultaneous labeling of three protein targets (i.e., TOMM20, histone 2B, and actin) in fixed cells, highlighting their utility for multicolor microscopy. MiniVIPER, TinyVIPER, and PunyVIPER are small and robust peptide tags for selective labeling of cellular proteins.

Introduction

Imaging by fluorescence microscopy (FM) is a powerful method for investigating how cellular proteins organize in space and time. Such studies typically use fluorescent labels to visualize selected proteins or cellular features. Organelles are often labeled using small-molecule fluorescent stains that function in relation to the organelle’s properties, such as membrane potential (mitochondria), DNA intercalation (nucleus), or pH (lysosomes).1 However, a shortage of protein-specific stains restricts them primarily to organelle imaging.2,3

In comparison, protein targets are usually identified by immunolabeling or fusion to a genetic tag. Immunolabeling has several major drawbacks. First, many antibodies lack specificity, resulting in dubious observations and poor reproducibility.4 A 2008 study found that 49% of antibodies failed to produce a staining pattern consistent with prior work or bioinformatics data.5 Second, immunolabeling is typically done post-fixation, precluding dynamic studies. Third, immunolabeling protocols can introduce cell damage and artifacts, again leading to misinformation.6

An excellent alternative to immunolabeling are genetic tags, where fluorescent labeling is achieved by the fusion of a protein tag to the protein of interest. There are many such tags for imaging in living cells.7,8 The most prominent example is the green fluorescent protein (FP).9 Although there are many differently colored FPs, they all have spectral properties determined (and limited) by a post-translationally generated chromophore.10 Two enzymes have also found widespread use as protein tags: a haloalkane dehalogenase (HaloTag)11 and a DNA alkyltransferase (SNAP-Tag).12 Enzyme tags accept fluorescent substrate mimics, which covalently bind within their active site. These tags confer an advantage over FPs since their substrates can be synthesized to exhibit optimal fluorescent properties (e.g., quantum yield, emission, and photostability). However, these and other protein tags13,14 are relatively large (14–30 kDa), which can disrupt folding, trafficking, binding, or function of the target protein.15

Peptide tags (<8 kDa) are less likely to impact the tagged protein’s physiological structure or function. An early example was the tetracysteine tag (<1 kDa),16 although issues with toxicity and background labeling have limited its widespread use. In contrast, coiled coils have emerged as a versatile and biocompatible motif for labeling cellular proteins.17,18 They are comprised of a short peptide tag (4.3–7 kDa) that forms an alpha-helical coiled-coil heterodimer with a labeled peptide (“probe peptide”). The specificity of coiled coils is determined by their peptide sequence, with interstrand salt bridges and a hydrophobic interface driving dimerization-mediated labeling.19 By separating the tag from the label, coiled coils maintain high labeling specificity while minimizing the impact on the target. Another advantage is the ability to easily change reporters using variably labeled probe peptides.20

The first example of the coiled-coil tag approach was reported by Matsuzaki and co-workers,21 who used an E3-K3 heterodimer22 to label membrane receptors for imaging. Since 2008, several other groups have used coiled-coil tags for cell imaging.18 Tamamura and co-workers have developed fluorogenic tags,23−25 including one for imaging within living cells.26 Some have developed probe peptides that mediate a proximity-induced reaction to ligate a fluorophore to a receptor.24,27−30 One approach (“Peptide-PAINT”) used transient coiled-coil binding for super-resolution microscopy.31

One potential limitation of coiled-coil tags is that labeling is restricted to tags accessible on the surface of living cells because the probe peptides are live cell membrane impermeant. This feature is ideal, however, for observing dynamic receptor interactions and trafficking.17,32 This limitation suggests that intracellular proteins must be labeled post-fixation. Alternatively, a few methods enable probe peptides to access intracellular targets in living cells (e.g., cell penetrating peptides or reversible membrane permeabilization).26 In our work, we labeled the mitochondria and nuclei in living cells using coiled coils through intracellular delivery of probe peptides by hollow gold nanoshells.33 We tracked the dynamic behavior of labeled structures for up to 48 h.

Since 2017, we have described three coiled-coil tags termed versatile interacting peptide (VIP) tags. VIP tags are small (4.3–6.2 kDa) and can be used for various imaging applications.20,32−37 In 2017, we reported our first tag for live cell imaging of receptors: VIP Y/Z.37 Next, we described VIPER, a tag formed by dimerization between a genetically encoded tag (CoilE) and a probe peptide (CoilR).32 We showed that VIPER could highlight organelles in fixed cells. We observed receptor endocytosis in live cells by “pulse-chase” labeling and time-lapse imaging. VIPER also illustrated that VIP tags enable labeling with reporters matched to the application, such as quantum dots (Qdots) for correlative light and electron microscopy (CLEM).32 Our CLEM studies included quantitative analysis of receptor labeling, by which we found that VIPER labeling was more efficient than immunolabeling. Most recently, we developed the MiniVIPER tag,20 which was inspired by a coiled-coil heterodimer first reported by Vinson and co-workers.38 MiniVIPER is a heterodimer between MiniE and MiniR. We showed that MiniVIPER could be used with either VIPER or VIP Y/Z for labeling two cellular proteins at once.20 However, those tags were not designed to be orthogonal to MiniVIPER, limiting the utility of VIP tags for multiprotein labeling.

As prior studies suggested that it would be feasible to create a set of orthogonal coiled-coil tags,39−41 the goal of our current work was to design a set of self-sorting peptide tags optimized for multiprotein labeling in cells. We aimed for this set of peptides to be biocompatible and bioorthogonal for stable, high-affinity labeling. To this end, we created two new tags, TinyVIPER and PunyVIPER, which were designed de novo to be used with MiniVIPER. We characterized each tag in vitro and found that the coiled-coil heterodimers interacted with low nanomolar affinity and high stability. In cells, each tag enabled selective labeling of proteins, including transferrin receptor 1 (TfR1), actin, TOMM20, and histone 2B (H2B). We show that VIP-tagged TfR1 retained physiological ligand binding and internalization patterns. Pairwise and three-way combinations of these tags showed bright, target-specific labeling, without cross-reactivity, of up to three distinct proteins in cells.

Materials and Experimental Details

See the Electronic Supporting Information (ESI) for a detailed description of all materials and methods used.

Results and Discussion

Design of New Self-Sorting VIP Tags

In the current work, we present a set of six peptides that self-sort to form coiled-coil heterodimeric tags for protein labeling without cross-reactivity (Figure 1). We de novo designed TinyVIPER and PunyVIPER to be orthogonal to our previously reported MiniVIPER tag.20 Each tag is 4.3 kDa, making them among the smallest tags reported to date. Peptide tag sequences, properties, and structural depictions (i.e., helical wheel diagrams) are reported in Figure 2.

Figure 1.

Figure 1

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Cellular protein labeling via orthogonal, self-sorting VIP tags. A set of three peptide tags, MiniVIPER (red), TinyVIPER (green), and PunyVIPER (blue), enable selective protein labeling with fluorescent reporters. Labeling is mediated by formation of coiled-coil heterodimers between each tag and a corresponding fluorescent probe peptide. Specificity is determined by favorable electrostatic interactions optimized for cognate heterodimers over mismatched pairs.

Figure 2.

Figure 2

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Design and biophysical properties of orthogonal VIP tags. (A) Peptide sequences reported using a single-letter amino acid code, displayed relative to the heptad motif in the header. Positively and negatively charged amino acids that define the electrostatic motif are labeled in blue and red, respectively. Asn (N) matches are bolded. Biophysical properties were determined by CD spectroscopy. The denaturant GdnHCl concentration where the coiled coil is 50% folded is reported as [GdnHCl]1/2. The free energy of heterodimer unfolding without the denaturant (ΔGH2O) was calculated by linear extrapolation, and the dissociation constant, KD(app), was calculated from ΔGH2O. (B–D) Helical wheel diagrams for each heterodimeric coiled coil. Residues are color-coded: charged = blue (+) or red (−); polar = orange; and hydrophobic = gray. Favorable salt bridges are illustrated by a blue dashed line. Diagrams were generated using DrawCoil1.049 (https://grigoryanlab.org/drawcoil/).

Each dimeric pair has characteristic alpha-helical coiled-coil interactions, including heptad repeats (denoted as abcdefg), salt bridges, and a hydrophobic interface. Each tag is four heptads, with charge distribution designed for interaction specificity and orthogonality. Residues at the a and d positions form a hydrophobic interface between the two coils that drives heterodimer formation, while interstrand salt bridges (g ↔ e′) enforce heterodimer specificity.42 We used a Glu–Arg electrostatic pair, which is 0.35 kcal/mol more stable than the Glu–Lys pair that is commonly used in other coiled-coil dimers.39,43,44 There are eight favorable salt bridges upon the formation of each optimized parallel heterodimer: MiniVIPER (Figure 2B), TinyVIPER (Figure 2C), or PunyVIPER (Figure 2D). TinyVIPER is a product of TREER–TinyERRE heterodimer formation, with the charge distribution indicated in the peptide name. PunyVIPER is a heterodimer of PuRRRE and PunyEEER.

Alpha-helical coiled coils can form parallel and anti-parallel dimers. We included an a position Asn–Asn match to promote formation of parallel dimers by hydrogen bonding between opposing coils.39,45−47 The Asn–Asn match was included in the second heptad of MiniVIPER, the third heptad for TinyVIPER, and the first and fourth for PunyVIPER. The remaining a position residues were Val, which further enhances the binding specificity because Val–Asn interactions are disfavored.46 Various potential homodimers and mismatched coiled-coil pairs are not expected to form due to destabilizing interactions,47,48 as illustrated in Figure S1 (see ESI).

Generation of Recombinant Peptides for Analysis and Imaging

Each peptide was made by recombinant expression in E. coli. Peptide sequences included a coil, followed by a short linker, reactive Cys, and a C-terminal hexahistidine tag (for purification). Cys was included to enable site-specific attachment of a fluorophore. In prior work, we made probe peptides using the standard thiol–maleimide conjugation32 (i.e., Tris buffer pH 7.2 with excess TCEP and reactive fluorophore) or Weiss’s solid-state labeling method.20,50 However, ligation was often inefficient. Unmodified and fluorophore-labeled peptides could not be separated, resulting in labeling reagents contaminated with up to 70% non-fluorescent peptides.20 For the current work, we optimized labeling by adapting a protocol published by Watts and co-workers.51 This method quenches the reducing agent (i.e., TCEP) before the addition of excess reactive fluorophore. After adopting this quenching step, we routinely achieved 50–90% labeling efficiencies (Table S4). We encountered an unexpected difficulty attaching a fluorophore to TREER. Therefore, TREER was used solely as the genetic tag for TinyVIPER labeling. Our method of making probe peptides uses commercially available fluorophores available in a range of colors and properties, making it feasible for others to rapidly generate probe peptides.

Biophysical Characterization of Coiled-Coil Heterodimers

We analyzed the secondary structure of the peptides and heterodimers by circular dichroism (CD) spectroscopy. We measured ellipticity between 200 and 260 nm for each peptide alone and for equimolar mixtures of cognate pairs (i.e., MiniE + MiniR, PunyEEER + PuRRRE, or TinyERRE + TREER). We did not evaluate non-cognate pairs. All monomer (Figure 3A) and heterodimer (Figure 3B) mixtures, except for PuRRRE, had curves with distinct minima at 208 and 222 nm indicative of an alpha-helical structure.52 In contrast, PuRRRE was unstructured. We assume that cognate pairs form alpha-helical heterodimers, although the high helicity of the monomers made our analysis inconclusive. We cannot rule out homodimer formation at the high peptide concentrations required for CD measurements, but the electrostatics would make such interactions disfavored (see Figure S1). We also saw no evidence of homodimer formation by FM (Figure S2), when probe peptides were used at experimentally relevant (nanomolar) concentrations.

Figure 3.

Figure 3

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MiniVIPER, TinyVIPER, and PunyVIPER form stable alpha-helical heterodimers. CD spectra of peptide monomers (A) and coiled-coil dimers (B). (C) Denaturation curves for MiniVIPER, TinyVIPER, and PunyVIPER. In (B,C), each peptide was present at a 1:1 molar ratio with 10 μM total peptide (5 μM each). Each curve in A–C represents three experimental replicates.

Next, we analyzed the conformational stability of heterodimers in increasing concentration of the denaturant guanidine hydrochloride (GdnHCl; 0 to 6 M) by measuring ellipticity at 222 nm. Figure 3C shows the fraction of the heterodimer folded, assuming that 100% was folded below 1 M GdnHCl and 0% was folded at 6 M GdnHCl. The denaturation curve suggested that all the three dimers had high stability, with each remaining 50% folded at ∼3.5 M GdnHCl. The observed ellipticity of each dimer with the denaturant was used to extrapolate the free energy of unfolding in the absence of the denaturant (ΔGH2O). All the three heterodimers were found to have similar ΔGH2O (∼11 kcal/mol) (Figure 2A).

The denaturation curves were used to calculate the apparent dissociation constant (KD(app)) using the method described by Litowski and Hodges.22 Each heterodimer had low nanomolar affinity (<25 nM) (Figure 2A). These values are consistent with the low nanomolar affinity measured for other four heptad coiled coils53 and match our prior prediction for MiniVIPER’s KD.20 The CD measurements required peptides to be at micromolar concentrations in order to be detected, raising the possibility that the true KD is quite a bit lower.54

Receptor Trafficking Observed with TinyVIPER and PunyVIPER

We established that each new VIP tag enabled specific labeling of proteins in living cells by imaging TfR1, a key component of the iron uptake machinery. We selected TfR1 as a model protein due to its well-characterized localization, trafficking, and ligand (transferrin) binding.55,56 Each VIP tag was introduced at the C-terminus, which is an extracellular domain for this receptor. Chinese hamster ovary (CHO) TRVb cells57 expressing tagged receptors were cooled to pause endocytosis before treatment with fluorescent transferrin (Tf-488) and 100 nM Sulfo-Cyanine 5 (Cy5)-labeled probe peptide. Cells were fixed immediately (0 min) or after receptor endocytosis (10 min at 37 °C) and then imaged by confocal FM.

We found that each VIP tag enabled selective labeling of TfR1 (Figure 4). Cells expressing VIP-tagged TfR1 showed a distinct signal at the cell surface (0 min) and in endocytic vesicles upon internalization (10 min). Cells expressing untagged TfR1 bound Tf-488 but not probe peptide. The signal from membrane-associated tagged TfR1 colocalized with Tf-488, as measured by Pearson’s correlation coefficient (PCC) (0 min = 0.60–0.83), and colocalization was maintained upon endocytosis (10 min = 0.47–0.63) (Figure S3). These colocalization metrics provide strong support that the VIP-tagged receptor retained the ligand-binding function. Furthermore, these observations suggest that VIP-tagged TfR1 retained physiological localization, trafficking, and function.

Figure 4.

Figure 4

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VIP-tagged TfR1 receptor localizes to the membrane, binds Tf, and traffics by endocytosis. CHO TRVb cells expressing VIP-tagged TfR1 were labeled live (4 °C) with 100 nM Cy5-conjugated probe peptide and 50 ng/mL Tf-488. Cells were fixed immediately after labeling (0 min) or after incubation at 37 °C to allow receptor–ligand internalization (10 min). (A) Cells expressing TfR1-PuRRRE or untagged TfR1 labeled with PunyEEER-Cy5 and Tf-488. (B) Cells expressing TfR1-PunyEEER or TfR1 labeled with PuRRRE-Cy5 and Tf-488. (C) Cells expressing TfR1-TREER or TfR1 labeled with TinyERRE-Cy5 and Tf-488. Micrographs are false-colored (Cy5, magenta; Tf-488, green; and Hoechst, blue), and green-magenta overlap appears white in the merge. The PCC between the Cy5 and 488 channels is reported in the lower left of each merge micrograph. Scale bars represent 20 μm.

We did not observe the Cy5 signal in untransfected cells or cells expressing untagged TfR1, illustrating that labeling was highly specific. We occasionally observed the probe peptide associated with cellular debris, as shown in the lower left panel in Figure 4C. For PunyVIPER (Figure 4A,B) and MiniVIPER20 (Figure S4), we demonstrated that either peptide could be encoded as the tag. We expect the same to be true of TinyVIPER, although we were limited to the use of TREER as the tag in the absence of fluorescent TREER probe peptide. Altogether, these results show that PuRRRE, PunyEEER, TREER, MiniE, and MiniR tags all enabled selective labeling of proteins in live cells.

Next, we looked at labeling specificity by treating cells expressing VIP-tagged TfR1 with each of the five Cy5-labeled probe peptides. Cells were selected based on the Tf-488 signal, and identical acquisition settings were used to image all cells expressing a given tag. We observed no homodimerization between each tag and the identical probe peptide (Figure S2). Surprisingly, TfR1-MiniE was labeled by both MiniR-Cy5 (cognate) and TinyERRE-Cy5 (non-cognate) probe peptides. Therefore, we did not use MiniE as a tag in multicolor labeling experiments. For receptors tagged with MiniR, PunyEEER, PuRRRE, and TREER, the brightest signal was observed for cells treated with the cognate probe peptide. We sought to detect any minimal labeling that might occur through non-cognate dimerization. To do this, we adjusted the displayed image contrast settings for the Cy5 signal by setting the highest pixel value 10-fold lower. We observed faint labeling for three non-cognate pairs (TfR1-PunyEEER + MiniE-Cy5, TfR1-PunyEEER + MiniR-Cy5, TfR1-TREER + MiniR-Cy5). The minimal cross-reactivity can be circumvented through experimental design, as shown below.

VIP-Mediated Two-Color Labeling of Organelles

After each tag was validated individually, we next used pairs of VIP tags to label two distinct targets in cells. We selected proteins with defined localization patterns, including TfR1 (cell membrane), actin (cytoskeleton), TOMM20 (mitochondria), and H2B (nucleus). First, we showed that TinyVIPER (actin) and PunyVIPER (mitochondria) could be used together to label cellular proteins without cross-reactivity (Figure 5). Human osteosarcoma cells (U-2 OS) expressing mEmerald FP (mEm)-TREER-actin and TOMM20-PuRRRE were fixed and treated with probe peptides (TinyERRE-Cy3 and PunyEEER-Cy5). PunyVIPER (Cy5) labeling was restricted to the mitochondria, which retained the normal morphology. TinyVIPER (Cy3) labeled actin, with the signal colocalized with green fluorescence (mEmerald), as expected. We did not observe the Cy5 signal in the cytoskeleton or the Cy3 signal in the mitochondria, suggesting that the two tags are orthogonal.

Figure 5.

Figure 5

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VIP tags enable specific labeling of two protein targets in cells. (A) U-2 OS cells expressing mEm-TREER-actin and TOMM20-PuRRRE were labeled post-fixation with PunyEEER-Cy5 and TinyERRE-Cy3 to enable simultaneous visualization of mitochondria and actin filaments. Micrographs are presented as a maximum intensity projection of a confocal z-stack. (B) CHO TRVb cells expressing TfR1-PunyEEER and H2B-MiniR-mEm were labeled with PuRRRE-Cy5 (live) and MiniE-Cy3 (post-fixation). Micrographs are false-colored (Cy3, magenta; Cy5, yellow; and mEm, green), and the merge includes Hoechst nuclear stain (blue). Scale bars represent 20 μm.

Next, we evaluated PunyVIPER and MiniVIPER for imaging H2B (nucleus) and TfR1 (cell membrane) in CHO TRVb cells (Figure 5B). In this example, we employed both live cell receptor labeling and fixed cell intracellular labeling. Cells expressing TfR1-PunyEEER and H2B-MiniR-mEm were first labeled with PuRRRE-Cy5 live to restrict labeling to the membrane-localized receptor. The cells were then fixed, treated with MiniE-Cy3, and imaged by confocal FM. Again, VIP-mediated labeling was specific, and tagged proteins retained their physiological organelle localization. MiniE-Cy3 colocalized with mEmerald within the nucleus.

In both dual-labeling experiments, there was no non-specific labeling in untransfected cells, cells expressing a single VIP-tagged protein, or cells transfected to express untagged proteins (Figure S5). Moreover, VIP tags did not appear to affect the normal localization of actin, TOMM20, H2B, or TfR1. Additional combinations of two VIP tags are included in Figure 6. Altogether, these studies illustrate that MiniVIPER, TinyVIPER, and PunyVIPER can be used to label and observe two protein targets simultaneously by FM.

Figure 6.

Figure 6

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MiniVIPER, TinyVIPER, and PunyVIPER enable simultaneous imaging of three distinct protein targets in fixed cells. (A) U-2 OS cells expressing TREER-actin, H2B-MiniR, and TOMM20-PuRRRE were fixed and then treated simultaneously with PunyEEER-Cy5, TinyERRE-Cy3, and MiniE-AF488. (B–D) Cells expressing two VIP-tagged proteins (as indicated) and labeled with all three probe peptides. Cells were imaged by confocal FM imaging, and micrographs represent maximum intensity projections from z-stacks. Micrographs are false-colored (Cy5, yellow; Cy3, magenta; AF488, green; and Hoechst, blue). Scale bar represents 20 μm.

Simultaneous VIP Labeling of Three Distinct Cellular Targets

Our objective herein was to create a set of bioorthogonal coiled-coil tags that expand the ability to label multiple proteins in cells. Therefore, we finally investigated whether TinyVIPER, PunyVIPER, and MiniVIPER could be used together to label three distinct cellular targets without cross-reactivity. We chose to label the cytoskeleton (actin), the nucleus (H2B), and the mitochondria (TOMM20) due to their distinct sub-cellular localizations. Fixed cells were labeled simultaneously with three spectrally distinct probe peptides (i.e., PunyEEER-Cy5, TinyERRE-Cy3, and MiniE-AF488) and then imaged (Figure 6). We observed selective labeling of each intended protein target by its corresponding probe peptide. Notably, there was no cross-labeling between mismatched pairs nor non-specific labeling in cells (Figure S6). These results provide direct evidence that MiniVIPER, TinyVIPER, and PunyVIPER are orthogonal peptide tags that can be used to simultaneously label three cellular targets.

Conclusions

As reported herein, the VIP tag repertoire has been expanded to include TinyVIPER and PunyVIPER for concurrent use with MiniVIPER. These coiled-coil heterodimers have high affinity (KD(app) < 25 nM) and stability, even under strongly denaturing conditions. These tags are ideal for labeling multiple cellular proteins at once due to several key characteristics. Foremost, they form a selective, bioorthogonal set of coiled-coil tags. The tags are small (4.3 kDa), making them unlikely to disrupt physiological protein functions, interactions, or localization. Our studies of VIP-tagged TfR1 support this assertion. Additionally, each tag is compatible with observing receptor trafficking in living cells. Last, VIP tags can be used to label proteins with diverse reporters, including fluorophores, biotin, and electron dense particles.20,32

These features highlight VIP tags as an advantageous alternative to conventional immunolabeling or large protein tags (e.g., FPs). Additionally, these features suggest their utility for other advanced imaging applications. The live cell compatibility of these tags would allow dynamic imaging of multiple cell receptors in parallel. As noted, live cell labeling is restricted to cell surface-exposed VIP tags. However, this limitation can be advantageous for labeling and tracking populations of receptors over time. We demonstrate this feature in Figure 4, where we use VIP tags to observe TfR1 endocytosis within 10 min of labeling.

For the current study, we showed that VIP tags could be inserted into proteins at the N-terminus, C-terminus, or intragenically. We assumed that a protein site that tolerates a FP (e.g., mEmerald) fusion would also accept a VIP tag. It is feasible that VIP tags are well tolerated because they mimic alpha-helical linkers found in nature.58 In all cases, we compared the behavior of untagged proteins with VIP-tagged proteins by imaging. For any new protein fusion, we recommend analyzing the localization, trafficking, binding, and function of the target protein before and after the introduction of a VIP tag. Further guidance on using genetic tags have been published.59,60

Beyond the studies presented here, VIP tags have the potential to advance labeling methodologies for cell-based EM and correlative fluorescence and EM, where labeling methods for target identification are currently limited. In future work, we plan to use VIP tags for multicolor CLEM studies of receptor organization. Overall, the VIP toolkit, including TinyVIPER, PunyVIPER, and MiniVIPER, offers many advantages for multiprotein investigations through imaging.

Acknowledgments

The authors are grateful to our colleagues at Oregon Health & Science University for their advice and guidance. Special appreciation is due to S. Kaech Petrie and the staff of the Advanced Light Microscopy Core, S. Reichow, and C. Enns. We appreciate the contributions of past lab members to the development of VIP tags.

Supporting Information Available

The Supporting Information is available free of charge at https://pubs.acs.org/doi/10.1021/acs.biochem.2c00712.

  • Detailed description of cloning, probe peptide generation, CD spectroscopy and analysis, and cell studies (e.g., transfection, labeling, imaging, and data processing) (PDF)

Accession Codes

TfR1: P02786; H2B: P06899; TOMM20: Q15388; and Actin: P60709.

Author Contributions

† A.S. and K.L.D. contributed equally to this work.

This research was supported by generous funding from Oregon Health & Science University and the National Institutes of Health (R01 GM122854).

The authors declare no competing financial interest.

Supplementary Material

bi2c00712_si_001.pdf (2.2MB, pdf)

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