Antibodies not only protect humans and other animals against disease-causing bacteria and viruses. They can also be used as tools for medical diagnostics and basic research. Conventional antibodies consist of light and heavy protein chains, and both are required to bind to target molecules (or antigens). Alpacas, llamas and camels, however, possess simpler antibodies that lack light chains and bind to antigens via a single protein domain. Such domains can be produced in "re-programmed" bacteria and are then called nanobodies. Compared to normal antibodies, nanobodies are 10-fold smaller, which is of great advantage in virtually all practical applications.

Pleiner et al. made nanobodies against the nuclear pore complex (or NPC for short) – a nanoscopic machine for transporting large biological molecules in and out of the cell’s nucleus. These nanobodies can be linked to dyes called fluorophores and then used to stain NPCs so that they can be observed under a microscope.

When fluorophores were attached, in the traditional way, via the amino acid lysine, all tested nanobodies performed poorly in fluorescence microscopy - pointing to a systematic problem. Pleiner et al. therefore explored an alternative, namely to label nanobodies via engineered cysteines. This was counterintuitive, because nanobodies contain already two other cysteines that must not be modified and that normally form a stabilizing “disulfide” bond. Pleiner et al. found, however, that the labeling reaction is absolutely specific for the engineered surface cysteines when it is performed at low temperature. This strategy consistently yielded imaging reagents that could effectively deliver fluorophores as close as 1-2 nanometers to their antigens. Nanobodies labeled in this way are therefore ideal to exploit the full potential of super-resolution microscopy.

The engineered surface cysteines proved also useful as "position sensors" to report which region of an antigen is actually contacted by a given nanobody.

Nanobodies are also used to purify protein complexes from crude cell extracts by a method called affinity chromatography. Previously, nanobodies were chemically attached to an insoluble matrix, and bound protein complexes were released under conditions that destroy interactions between proteins. Pleiner et al. now replaced the destructive step with a step that uses an enzyme to cut a bond and gently detach the nanobody (along with any bound protein complex) from the matrix. Bound protein complexes thus stay intact and can be studied further. In the future, this strategy can be applied to nanobodies that recognize tags commonly added to proteins (i.e. GFP) to isolate virtually any protein complex for functional assays or structural analyses.

Nanobodies represent antigen-binding domains of 'heavy-chain-only' camelid antibodies and are typically selected by phage display from an immune library (Hamers-Casterman et al., 1993; Arbabi Ghahroudi et al., 1997; Muyldermans, 2013). Their small size (~13 kDa), monoclonal nature and high specificity are ideal for applications like affinity purification or protein detection and localization (Helma et al., 2015). Their utility as crystallization chaperones is also widely appreciated (Pardon et al., 2014; Desmyter et al., 2015).

Nanobodies are commonly expressed in Escherichia coli and secreted into the oxidative periplasm, where their conserved internal disulfide bond can be formed (Vincke et al., 2012; Pardon et al., 2014; Fridy et al., 2014). Periplasmic expression comes, however, with several drawbacks. For example, it often results in low final yield (Baneyx and Mujacic, 2004), probably due to saturation of the secretion machinery and aggregation of precursor proteins in the cytoplasm. The limited set of chaperones and high proteolytic activity in the periplasm also restrict the choices of fusion tags that can be used (de Marco, 2009; Feilmeier et al., 2000). Furthermore, the purification of periplasmic proteins involves considerably more hands-on time than purification from the cytoplasm.

In typical affinity chromatography applications, nanobodies are covalently attached to a resin, which later necessitates harsh conditions for the elution of bound target proteins (Rothbauer et al., 2008; Fridy et al., 2014). This is appropriate for an identification, but hardly for any further downstream structural or functional analysis of the purified target proteins. As a workaround, a native isolation of protein A-tagged protein complexes using a specific nanobody modified with a dithiothreitol (DTT)-cleavable crosslinker was recently reported (Fridy et al., 2015). However, the achievable yield was modest, as most of the isolated complexes resisted release. Furthermore, the presence of any thiol-reducing agent (like DTT or glutathion) during binding is incompatible with this method.

In traditional indirect immunofluorescence, epitopes are initially decorated with a primary antibody and detected with a fluorophore-labeled secondary one, each around 12–15 nm in size (Harris et al., 1998). The effective displacement between label and epitope can reach up to 24–30 nm and thus significantly deteriorate the achievable precision and accuracy of protein localization by super-resolution fluorescence microscopy (Hell, 2009; Huang et al., 2009). Nanobodies (diameter: 4 nm) are an ideal solution to this problem (Ries et al., 2012; Szymborska et al., 2013). This, however, requires a direct nanobody labeling. Ideally, labeling should be site-specific, so that the remaining small displacement between epitope and fluorescent dye can be predicted and corrected for in the measurements.

So far, nanobodies were labeled at lysines by N-hydroxysuccinimide (NHS) ester fluorophores (Ries et al., 2012; Fridy et al., 2014), which is random and rarely quantitative. As we show below, it also deteriorates signal-to-background ratios or even completely abolishes epitope recognition. A workaround to this basic problem was the addition of a C-terminal oligo-lysine stretch to divert labeling from nanobody framework lysine residues (described for the anti-GFP nanobody 'Enhancer' in Platonova et al., 2015). This, however, increases the epitope-label distance again. Furthermore, fluorescent labeling of nanobodies using Sortase A was presented (Witte et al., 2012). This strategy is limited to the N- or C-terminus and uses modified fluorophores that are not readily available. Adding an extra C-terminal cysteine (for subsequent maleimide modification) to a periplasmically expressed nanobody was also not a satisfying solution, because it led to a severe reduction in yield and caused extensive dimerization (Massa et al., 2014).

Hence, we explored solutions to the above-described limitations of the current nanobody technology. We demonstrate functional cytoplasmic expression of nanobodies with protease-cleavable tags for native affinity purification and with engineered cysteines for site-specific fluorescent labeling. We chose the Xenopus nuclear pore complex (NPC) as a model target and developed a toolbox of high-affinity nanobodies against its major constituent proteins, nucleoporins (Nups), which occur in large subcomplexes. Using specific nanobodies, we purified their target protein complexes from Xenopus egg extract in a single step with native elution based on proteolytic matrix-release. This allowed a direct analysis of nanobody-purified endogenous Nup complexes by negative stain electron microscopy. Labeling these anti-Nup nanobodies with NHS ester fluorescent dyes for imaging often produced non-functional reagent or significant background staining. We therefore implemented a simple and generally applicable strategy for obtaining site-specifically fluorophore-labeled nanobodies of superior imaging quality. It involves engineered cysteines at the nanobody surface, their modification with maleimide fluorophores, and leaves the internal framework cysteines fully intact. This strategy allowed super-resolution imaging of NPCs with a negligible label displacement and very low background. A novel strategy for rapid mapping of conformational nanobody epitopes via crosslinking mass spectrometry involving the engineered surface cysteines is also presented here.

NPCs are gateways for nucleocytoplasmic transport. Their very large size of ≈110 MDa not only places them amongst the largest molecular machines, but also poses formidable challenges for any structural and functional investigation (reviewed in Hurt and Beck, 2015). Nucleoporins are organized in multiple subcomplexes around a central eightfold rotational symmetry axis. Certainly, the most characteristic subcomplex is the Nup107-Nup160 or Y-shaped complex (Siniossoglou et al., 2000; Vasu et al., 2001). Its essential role in NPC assembly (Harel et al., 2003; Walther et al., 2003) as well as its structural organization and relative position within the NPC have been studied intensely (Bui et al., 2013; Eibauer et al., 2015; Kelley et al., 2015; Stuwe et al., 2015b; von Appen et al., 2015). The inner ring of the NPC scaffold comprises also essential Nup93-containing subcomplexes (Sachdev et al., 2012; Vollmer and Antonin, 2014), whose stoichiometry and orientation are less understood. Nup93 further anchors the trimeric Nup62•Nup58•Nup54 complex in the central transport channel (Finlay et al., 1991; Hu et al., 1996; Chug et al., 2015; Stuwe et al., 2015a).

Functional cytoplasmic expression of anti-NPC nanobodies

In order to provide new tools for studying NPCs, we generated nanobodies against constituents of the Xenopus NPC scaffold, namely Nup85, Nup93, and Nup155, as well as against Nup98 and the Nup62•Nup58•Nup54 complex. The latter two species were included because their Phe-Gly (FG)-repeat domains form a permeability barrier within the central NPC channel (Hülsmann et al., 2012). High-affinity nanobodies against all these targets were readily obtained from alpaca immune libraries by phage display.

We noticed that these nanobodies could be produced in the cytoplasm of various E. coli strains, as recently reported for other nanobodies (Olichon and Surrey, 2007; Zarschler et al., 2013; Djender et al., 2014). We also observed that fusing nanobodies behind a His₁₄-bdNEDD8 module (Frey and Görlich, 2014) increased their yield dramatically (Figure 1a).

Figure 1

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In order to test whether disulfide bond formation can occur in the cytoplasm and if this is important for nanobody function, we expressed an anti-Nup93 and an anti-Nup98 nanobody either in the periplasm, the reductive cytoplasm of E. coli BLR or in the cytoplasm of E. coli SHuffle. The latter strain contains a cytoplasmic disulfide isomerase and harbors mutations that render its cytoplasm (moderately) oxidative (Lobstein et al., 2012). The obtained nanobody variants were then treated with biotin-PEG₂₃-maleimide under denaturing conditions. Reduced nanobodies are thereby modified at their free cysteines, and the resulting size shift distinguishes them from disulfide-containing nanobodies (Figure 1b). While periplasmic secretion resulted in fully oxidized nanobodies, only a fraction of the SHuffle-expressed nanobodies contained a disulfide bond. Cytoplasmic expression in E. coli BLR yielded completely reduced nanobodies.

One could assume that the antigen affinity of nanobodies is negatively affected by a loss of their scaffold disulfide bond. A competition for antigen-binding revealed, however, no affinity difference between reduced and disulfide bond-containing anti-Nup98 nanobodies (Figure 1c). As expected, we observed by differential scanning fluorimetry (Niesen et al., 2007) a decreased thermostability of fully reduced anti-Nup98 and anti-Nup93 nanobodies (Figure 1d). Their melting temperatures of 47°C and 57°C are, however, still well above any reasonable incubation temperature for downstream applications.

All nanobodies that we obtained via phage display against a wide range of antigens could be functionally produced in the E. coli cytoplasm. Only very few of those nanobodies contained a second pair of cysteines that can form an additional, solvent-exposed disulfide bond between the antigen-binding loops CDR II and CDR III which likely contributes to the overall affinity (Govaert et al., 2012). However, most biochemical applications as well as imaging techniques like STORM require reducing conditions that disrupt accessible disulfide bonds, making such nanobodies a poor option anyway.

Native purification of endogenous protein complexes using nanobodies

Cytoplasmic expression of nanobodies provides a number of advantages. First, the yield often exceeds 100 mg per liter of culture and can be up to 30 times higher as compared to periplasmic expression (Figure 1a). Second, it saves hands-on time, because a cumbersome preparation of a periplasmic fraction is bypassed, and third, a far broader range of fusion modules can be used.

We exploited this for affinity purification of endogenous target protein complexes with nanobodies and native elution. For this strategy, we produced His₁₄-Avi-(GlySer)₉-SUMOStar-(GlySer)₉-nanobody fusions and purified them by Ni²⁺ chelate affinity chromatography and imidazole elution (Figure 2—figure supplement 1a). The Avi-tag can be biotinylated by cytoplasmic co-expression of the biotin ligase BirA in E. coli (Schatz, 1993; Beckett et al., 1999) or in vitro using the purified enzyme (Fairhead and Howarth, 2015). It then mediates binding of the purified nanobody to streptavidin magnetic beads. The interspersed long unfolded Gly-Ser spacers minimize steric hindrance effects. The SUMOStar module is an engineered SUMO variant that cannot be cleaved by endogenous eukaryotic desumoylases but by an engineered SUMOStar protease (LifeSensors), (Liu et al., 2008). In combination, these modules allow native elution of nanobody-bound target proteins or protein complexes by cleaving the tag with nanomolar concentrations of the SUMOStar protease. This strategy also provides a purer and more specific end product, because any protein species, which sticks non-specifically to the beads, will not be released. Thus, such highly specific protease elution makes the otherwise crucial control for matrix background-binding (Marcon et al., 2015; Mellacheruvu et al., 2013) essentially dispensable.

As a proof of principle, we purified five nucleoporin complexes from a Xenopus egg extract to near homogeneity (Figure 2a and b). For each complex we achieved a ≈10 000-fold enrichment within a single native purification step and yields of around 50%. The anti-Nup85 nanobody retrieved the ≈750 kDa nine-membered Y-complex as well as Tpr and Elys as specific but sub-stoichiometric binding partners. We obtained substantial amounts of the complex, namely 50–100 µg from as little as 2 ml egg extract, which initially contained ≈150 µg or ≈100 nM of the complex (Wühr et al., 2014). Post-elution with SDS sample buffer indicated a quantitative proteolytic release of the complex from the beads (Figure 2—figure supplement 1b).

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The anti-Nup155 nanobody retrieved Nup155 as a single species. This might appear surprising as Nup155 is thought to contact Nup93 and Nup53/35 within the inner ring of the NPC scaffold (Hawryluk-Gara et al., 2005; Hawryluk-Gara et al., 2008; Sachdev et al., 2012). We therefore assume that mitotic post-translational modifications transiently suppress interactions between these proteins. We also purified the Nup98•Gle2 and the Nup62•Nup58•Nup54 complex using anti-Nup98 and anti-Nup54 nanobodies, respectively. Here, we included a RanQ69L•GTP wash to release nuclear transport receptors, which otherwise would remain bound to the FG domains of Nup98 or the Nup62 complex (Figure 2—figure supplement 1c).

The anti-Nup93 nanobody purified the expected mixture of the two paralogous Nup93•Nup188 and Nup93•Nup205 complexes (Theerthagiri et al., 2010), which are also a part of the structurally least understood NPC inner ring. In this case, we analyzed the natively eluted material straightaway by negative stain electron microscopy (Figure 3a). Class averaging revealed characteristically curved α-solenoid fold-like particles, which are known to exhibit conformational flexibility (Figure 3b). The obtained structures were very reminiscent of the hook and eye-shaped structures reported earlier for the Nup188 and Nup205 orthologues from Saccharomyces cerevisiae (Amlacher et al., 2011) and Myceliophthora thermophila (Andersen et al., 2013). This suggests not only that the overall shape of the Nup93 complexes is conserved from fungi to vertebrates, but also that our single-step purification strategy for large protein complexes yields material of sufficient quality for a direct structural analysis.

Figure 3

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Site-specific fluorescent labeling of nanobodies

In order to use anti-Nup nanobodies to image their targets within intact NPCs, we initially modified them with NHS ester fluorophores. We found, however, that such NHS-labeled nanobodies performed remarkably poorly, in particular when far-red fluorophores were used. As documented by the specific examples below, none of the NHS-labeled nanobodies had sufficient probe quality to allow acquisition of STORM images. We therefore explored alternative and more reliable nanobody-labeling strategies.

One possibility was to label nanobodies at engineered (and reduced) cysteines with maleimides. This, however, posed the risk of modifying also the scaffold cysteines of the IgG-fold, which inevitably would cause an irreversible unfolding of the nanobodies.

To address this issue, we incubated reduced nanobodies with biotin-PEG₂₃-maleimide (Figure 4). After unfolding by urea, the scaffold cysteines became modified at either 37°C, 23°C, or 0°C. In native buffer, however, modification was quantitative only at 37°C, pointing to a transient exposure of the otherwise buried scaffold cysteines ('thermal breathing'). Importantly, they remained fully protected at 0°C, predicting that maleimide-labeling on ice would be fully selective for engineered surface cysteines.

Figure 4

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In order to better guide cysteine placement in the nanobody framework, we solved the crystal structure of the anti-Nup98 nanobody TP377 in complex with the globular Nup98 NPC anchor domain (residues 716–866) at 1.9 Å resolution (Figure 5a, Table 1). TP377 contacts its target through all three CDR loops and does not block the absolute Nup98 C-terminus, which anchors Nup98 via Nup96 or Nup88 to the NPC scaffold (Hodel et al., 2002; Griffis et al., 2003; Stuwe et al., 2012). The internal disulfide bond-forming cysteines Cys22 and Cys96 of TP377 are reduced in the crystal structure.

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Table 1

	Nup98•Nb TP377 complexa
Data collection
Space group	P41
Cell dimensions
a, b, c (Å)	66.59, 66.59, 87.90
α, β, γ (°)	90.00, 90.00, 90.00
Resolution (Å)	47.00-1.90 (1.95-1.90)b
Rsym or Rmerge	0.128 (>1)b
I / σI	27.7 (2.6)b
Completeness (%)	99.5 (98.7)b
Redundancy	27.4 (27.4)b
Refinement
Resolution (Å)	47.00-1.90
No. reflections
Measured	823105
Unique	30218
Rwork/Rfree	0.167 / 0.196
No. atoms
Protein	2176
Water	145
Wilson B-factor (Å2)	27.4
R.m.s. deviations
Bond lengths (Å)	0.010
Bond angles (°)	1.08
Ramachandran statistics (%)
Favored	98.0
Allowed	2.0
Outliers	-

1. ^aA single crystal was used for data collection.

2. ^bValues in parentheses are for highest resolution shell.

We next mutated solvent-exposed small residues (Gly, Ser, and Ala) at six alternative positions of the nanobody scaffold to cysteines (Figure 5b; Figure 5—figure supplement 1a). We found that not only all individual mutants, but also nanobodies with up to three ectopic surface cysteines were well expressed and highly soluble in E. coli (Figure 5—figure supplement 1b). Moreover, cysteines on all six positions on our model anti-Nup98 nanobody TP377 could be quantitatively labeled with maleimide fluorescent dyes (Figure 5c). Even nanobodies carrying three fluorophores were readily obtained. Importantly, competitive binding assays indicated that the introduction of ectopic cysteines and their modification did not impair binding to the Nup98 target (Figure 5—figure supplement 1c). Based on the crystal structure, we estimate that fluorophores attached via an N-terminal cysteine or A75C to anti-Nup98 nanobody TP377 can be as close as ~2 nm to the target Nup98.

Immunofluorescence with site-specifically labeled nanobodies

In order to test nanobodies in imaging, we grew Xenopus laevis XL177 cells on coverslips, digitonin-permeabilized their plasma membranes, incubated them with low nanomolar concentrations (1–10 nM) of labeled nanobody, and fixed them after several washing steps. In this workflow, even nanobodies with fixation-sensitive epitopes could bind their target.

We first tested anti-Nup98 nanobody TP377 carrying a single Alexa Fluor 647 maleimide at the six alternative positions (Figure 5d). In confocal laser scanning microscopy, all variants produced a very bright punctuate nuclear rim staining of XL177 cells, characteristic for NPCs, against a very low background.

Combining minimal label displacement with ease of cloning, we routinely labeled our nanobodies via an N-terminal cysteine. This way, all chosen NPC targets (Nup98, Nup93, Nup85 and Nup155) could be visualized with specific nanobodies carrying a single N-terminal Alexa Fluor 647 maleimide (Figure 6a). Despite the presence of only one dye molecule per nanobody, we again obtained very bright nuclear rim stains with very low background. Staining of Nup155 required a prior permeabilization with Triton X-100, probably because it is located in close proximity to the pore membrane and is likely buried by other NPC scaffold components (Eisenhardt et al., 2014; Mitchell et al., 2010).

Figure 6

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For a direct comparison of the NHS chemistry for nanobody-labeling at lysines with maleimide-labeling at engineered surface cysteines, we chose Alexa Fluor 647 as a fluorophore and the widely used anti-GFP nanobody Enhancer (Kirchhofer et al., 2010) as an example (Figure 6b). When a HeLa Nup153-GFP cell line was stained, we observed a brilliant NPC signal for the Alexa Fluor 647 maleimide-labeled 'Enhancer', which perfectly coincided with the (weaker) GFP signal, and an extremely low background (Figure 6c). In contrast, when this nanobody was labeled at lysines with Alexa Fluor 647 NHS ester, it produced strong nucleoplasmic and cytoplasmic background staining, which essentially obscured the specific signal. The degree of labeling (DOL) was the same for both variants.

When the Alexa Fluor 647 NHS-labeled 'Enhancer' was applied to XL177 cells (which lack a GFP-target), we again observed very strong background (Figure 6d). In contrast, its Alexa Fluor 647 maleimide-labeled counterpart behaved like a perfect negative control. High background-staining was also observed with the Alexa Fluor 647 NHS-labeled anti-Nup98 nanobody TP377. The anti-Nup93 nanobody TP179 contains a lysine in CDR II and even lost antigen-binding after NHS modification. In contrast, the Alexa Fluor 647 maleimide-labeled anti-Nup98 and anti-Nup93 nanobodies behaved as perfect imaging reagents and gave crisp NPC signals against very low backgrounds. This comparison indicated that modification of (multiple) framework lysines likely creates hydrophobic patches that favor unspecific binding and aggregation. This is certainly sequence context-dependent and milder when reducing the labeling density. However, we did not observe any such complications when labeling nanobodies via engineered cysteines.

Super-resolution imaging with site-specifically labeled nanobodies

Due to a diameter well below the diffraction limit, NPCs have been studied by super-resolution microscopy using either indirect immunofluorescence (Löschberger et al., 2012; Göttfert et al., 2013) or the anti-GFP nanobody (Szymborska et al., 2013). Site-specific fluorescent labeling of nanobodies via cysteines now reliably yields 'renewable' high-quality imaging reagents that can bring fluorophores very close to their target. We therefore tested the performance of our anti-Nup nanobodies in STORM imaging (Rust et al., 2006) of XL177 cell NPCs (Figure 7a–c).

Interestingly, singly Alexa Fluor 647 maleimide-labeled anti-Nup nanobodies were sufficient to produce enough localizations to reconstruct very detailed views of individual NPCs, where multiple copies of the imaged target proteins (Nup98, Nup93, and Nup155) appear arranged around the central NPC channel (Figure 7c). A whole nucleus stained with the model anti-Nup98 nanobody TP377 is shown in Figure 7a and magnified views of the nuclear envelope stained with anti-Nup93 and anti-Nup155 nanobody are shown in Figure 7b. Even after applying higher concentrations (~100–300 nM) of nanobody to saturate binding sites, we achieved very low background binding, indicating well-behaved imaging reagents.

Figure 7

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Rapid mapping of conformational epitopes via crosslinking mass spectrometry

Site-specifically labeled nanobodies enabled us to visualize their targeted epitope with high precision. Mapping the corresponding 'visible' epitopes would therefore reveal surface areas of the target that are accessible in a cellular environment. The complementary 'invisible' epitopes on the other hand, would hint to regions that are buried in interaction interfaces.

Epitope-mapping strategies based on binding assays to truncated or mutated antigens, co-crystallization or NMR observation of chemical shift perturbations are, however, not suited for high-throughput analysis or hardly applicable to conformational epitopes on protein complexes. We therefore considered crosslinking nanobodies to their target followed by sequencing of the crosslinked target peptide via mass spectrometry as a rapid epitope mapping strategy. Here, a crucial aspect is that a shorter crosslinker will provide a better spatial resolution, provided crosslinkable groups are in reach. As cysteines are by far the best crosslinkable groups, and because we had already placed cysteines at the nanobody surface in close proximity to bound targets, we assessed their suitability for epitope-mapping.

As a proof of principle, we crosslinked two anti-Nup93 nanobodies, with or without an N-terminal cysteine, to Nup93 using either an NH₂-to-NH₂ (Bis-NHS; BS3; 11.4Å) or an SH-to-NH₂ (Mal-NHS; BMPS; 5.9 Å) crosslinker (Figure 8a). For both anti-Nup93 nanobodies (TP179 and TP324), exclusive amine-crosslinking was very inefficient and produced only few nanobody•Nup93 adducts that run at higher molecular weight in SDS–PAGE. However, combining the N-terminal cysteine on the nanobody with the (far shorter) heterobifunctional crosslinker, produced very prominent nanobody•Nup93 crosslinks. Their position was then clearly identifiable by LC-MS/MS (Figure 8b–c, Figure 8—figure supplement 1).

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For a better visualization of the positions of the identified nanobody crosslinks we generated a structural model of Nup93^168-end using I-TASSER (Zhang, 2008), based on structures of its yeast ortholog (Jeudy and Schwartz, 2007; Schrader et al., 2008) (Figure 8c).

We used the anti-Nup93 nanobody TP179 in STORM imaging of Nup93 within the NPC and could now map its accessible epitope. TP179 binds to the middle region of the J-shaped structure of Nup93 surrounding residues K607 and K612 (Figure 8—figure supplement 1a–b), while TP324 has a C-terminal epitope surrounding lysines K762, K765, and K782 of Nup93 (Figure 8—figure supplement 1c–e). The C-terminal region of Nup93 was previously shown to be essential for NPC assembly (Sachdev et al., 2012). Accordingly, anti-Nup93 nanobody TP324 that targets the C-terminus of Nup93 does not stain intact NPCs (data not shown), but rather represents a good candidate to selectively disrupt NPC assembly.

We developed a well-characterized toolset of high-affinity nanobodies against the vertebrate NPC and established novel strategies to use these nanobodies to natively purify large NPC subcomplexes and to reliably label them with fluorophores for precise super-resolution localization. While these nanobodies will be very valuable to the nucleocytoplasmic transport field, we expect the presented strategies to be widely applicable to all nanobodies.

Nanobodies against a single epitope of a larger protein complex now allow a native single-step purification of the entire complex, and thus a subsequent structural and functional analysis. This is certainly especially useful for complexes that are not directly accessible to recombinant production. Furthermore, nanobody-purified endogenous complexes can be used as antigens for another round of immunization, and binders to all complex components can then be selected from the successive nanobody library. Mapping epitopes via crosslinking mass spectrometry will become especially important when selecting nanobodies against such complex antigens (like large subcomplexes, whole organelles or vesicles) that cannot be made recombinantly. Combining these strategies therefore has the potential to significantly increase throughput in the selection and identification of renewable binders to eukaryotic proteomes (Colwill et al., 2011).

Finally, we introduced a method for a reliable fluorescent labeling of nanobodies using surface cysteines and maleimide chemistry. This way, we obtained well-behaved imaging reagents that can bring fluorescent dyes as close as 1–2 nm to their target. Maleimide-labeled nanobodies consistently recognized their antigens far better and produced less background than the corresponding NHS-modified variants. NHS esters have the additional disadvantage that they react not only with amino groups, but also rapidly hydrolyze in aqueous buffers. This makes it difficult to adjust labeling densities and requires adding them in substantial molar excess. In contrast, maleimide-labeling of exposed cysteines is quantitative even with just stoichiometric amounts of labeling reagent and thus far more economical.

Site-specific and quantitative fluorescent labeling of nanobodies is going to be crucial for super-resolution microscopy aiming at a detailed structural analysis or determination of absolute protein copy numbers. It also allows predicting the effective label displacement, a fact that will be especially important when applying particle averaging techniques to localization microscopy data (precision of <1 nm reported by Szymborska et al., 2013). Because of its well-defined dimension and symmetric structure, the NPC has become a benchmark for many new advancements of super-resolution microscopy (Schermelleh et al., 2008; Szymborska et al., 2013; Göttfert et al., 2013). The anti-NPC nanobodies described here excelled in super-resolution imaging; they can be renewably produced in high yields and are therefore ideal labeling reagents for such benchmark studies.

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Author details

1. Tino Pleiner

   Department of Cellular Logistics, Max Planck Institute for Biophysical Chemistry, Göttingen, Germany

   Contribution

   TP, Conception and design, Acquisition of data, Analysis and interpretation of data, Drafting or revising the article

   Competing interests

   The authors declare that no competing interests exist.

   "This ORCID iD identifies the author of this article:" 0000-0002-5104-0315

2. Mark Bates

   Department of NanoBiophotonics, Max Planck Institute for Biophysical Chemistry, Göttingen, Germany

   Contribution

   MBa, Acquisition of data, Analysis and interpretation of data

   Competing interests

   The authors declare that no competing interests exist.

   "This ORCID iD identifies the author of this article:" 0000-0003-0668-5277

3. Sergei Trakhanov

   Department of Cellular Logistics, Max Planck Institute for Biophysical Chemistry, Göttingen, Germany

   Contribution

   ST, Acquisition of data, Analysis and interpretation of data

   Competing interests

   The authors declare that no competing interests exist.

   "This ORCID iD identifies the author of this article:" 0000-0002-1326-6153

4. Chung-Tien Lee

   1. Bioanalytical Mass Spectrometry, Max Planck Institute for Biophysical Chemistry, Göttingen, Germany
   2. Bioanalytics, Institute for Clinical Chemistry, University Medical Center Göttingen, Göttingen, Germany

   Contribution

   C-TL, Acquisition of data, Analysis and interpretation of data

   Competing interests

   The authors declare that no competing interests exist.

5. Jan Erik Schliep

   3D Electron Cryo-Microscopy Group, Max Planck Institute for Biophysical Chemistry, Göttingen, Germany

   Contribution

   JES, Acquisition of data, Analysis and interpretation of data

   Competing interests

   The authors declare that no competing interests exist.

   "This ORCID iD identifies the author of this article:" 0000-0001-7103-5954

6. Hema Chug

   Department of Cellular Logistics, Max Planck Institute for Biophysical Chemistry, Göttingen, Germany

   Contribution

   HC, Drafting or revising the article, Contributed unpublished essential data or reagents

   Competing interests

   The authors declare that no competing interests exist.

   "This ORCID iD identifies the author of this article:" 0000-0002-9889-7076

7. Marc Böhning

   Department of Cellular Logistics, Max Planck Institute for Biophysical Chemistry, Göttingen, Germany

   Contribution

   MBö, Acquisition of data, Analysis and interpretation of data

   Competing interests

   The authors declare that no competing interests exist.

   "This ORCID iD identifies the author of this article:" 0000-0002-4245-5113

8. Holger Stark

   3D Electron Cryo-Microscopy Group, Max Planck Institute for Biophysical Chemistry, Göttingen, Germany

   Contribution

   HS, Analysis and interpretation of data, Drafting or revising the article

   Competing interests

   The authors declare that no competing interests exist.

9. Henning Urlaub

   1. Bioanalytical Mass Spectrometry, Max Planck Institute for Biophysical Chemistry, Göttingen, Germany
   2. Bioanalytics, Institute for Clinical Chemistry, University Medical Center Göttingen, Göttingen, Germany

   Contribution

   HU, Analysis and interpretation of data, Drafting or revising the article

   Competing interests

   The authors declare that no competing interests exist.

   "This ORCID iD identifies the author of this article:" 0000-0003-1837-5233

10. Dirk Görlich

    Department of Cellular Logistics, Max Planck Institute for Biophysical Chemistry, Göttingen, Germany

    Contribution

    DG, Conception and design, Analysis and interpretation of data, Drafting or revising the article

    For correspondence

    goerlich@mpibpc.mpg.de

    Competing interests

    The authors declare that no competing interests exist.

    "This ORCID iD identifies the author of this article:" 0000-0002-4343-5210

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