Prior to the recent determination of the TAPBPR-MHC I crystal structures (Jiang et al., 2017; Thomas and Tampé, 2017), we docked our model of TAPBPR onto a previously determined structure of HLA-A2, using our mutagenesis data that identified critical regions in the TAPBPR-MHC I interface (Hermann et al., 2013). Our docking identified a region of TAPBPR that lies close to the peptide binding groove of MHC I, in the proximity of the F pocket (Figure 1a, dotted circle). This region contained a loop that differs between tapasin and TAPBPR. In tapasin, this loop appears to be rather short and is not sufficiently well ordered in the crystal structure to be visible (Dong et al., 2009), while in TAPBPR this loop is significantly longer (Figure 1b). The two crystal structures of the TAPBPR-MHC I complex support our prediction regarding the arrangement of TAPBPR relative to MHC I, including this loop region being very near the F pocket (Figure 1c). However, the position and orientation of the loop in the structures is poorly defined. In the structure from Jiang et al. (2017), this loop is not sufficiently well ordered to be modelled. In the structure from Thomas and Tampé (2017), the loop has been modelled; however, the electron density into which it was built is not well defined: several side chains and even several of the backbone atoms do not fit the electron density well (Figure 1d). It is likely that alternative orientations of the loop would also satisfy the crystallographic data presented. Therefore, it is critically important to verify whether there is any functional role of this loop in peptide editing.

Figure 1

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To test whether the K22-D35 loop of TAPBPR was a functionally important region for mediating peptide selection, we first replaced all the residues in the loop with either glycine, alanine or serine, to produce a TAPBPR variant with a potentially functionless loop (TAPBPR^Øloop) (Table 1). More subtle mutations of this loop were designed based on the work by Springer and colleagues, who demonstrated that dipeptides carrying long hydrophobic residues are able to bind to the peptide binding groove of recombinant MHC I and enhance peptide dissociation (Saini et al., 2015). Thus, we explored whether a leucine residue at position 30 of the mature TAPBPR protein, the only long hydrophobic residue within the entire loop, was involved in peptide exchange on MHC I. We created two TAPBPR variants in order to test this. First, we replaced the leucine with glycine in the TAPBPR^WT molecule (TAPBPR^L30G). Second, we reintroduced leucine 30 into TAPBPR with the dysfunctional loop (TAPBPR^ØG30L) (Table 1). Upon transduction into TAPBPR-deficient HeLaM cells (HeLaM-TAPBPR^KO), steady state expression of all the TAPBPR loop mutants was similar to TAPBPR^WT and all variants interacted equally well with both MHC I and UGT1 (Figure 1e), suggesting the overall protein stability and structure of TAPBPR was not significantly affected by the changes to the loop.

Recently, we have developed two novel assays which can be used to measure TAPBPR-mediated peptide exchange on MHC I molecules. The first assay established takes advantage of the small proportion of TAPBPR that escapes to the cell surface upon its over-expression in cell lines (Ilca et al., 2018). We have demonstrated that plasma membrane expressed wild-type TAPBPR efficiently mediates peptide exchange on cell surface HLA-A*68:02 molecules found on HeLaM cells (Ilca et al., 2018). We have also previously shown that TAPBPR-mediated peptide exchange in this assay occurs directly on the cell surface, given that it works on cells incubated at 4°C, which inhibits membrane trafficking (Ilca et al., 2018). To initially explore the functional importance of the K22-D35 loop, we tested whether plasma membrane expressed TAPBPR^Øloop was capable of mediating peptide exchange on HLA-A*68:02 to a similar extent as TAPBPR^WT. When exploring the ability of TAPBPR to promote peptide association, we found plasma membrane expressed TAPBPR^Øloop enhanced the binding of an exogenous fluorescent peptide specific for HLA-A*68:02 (YVVPFVAK*V) onto cells to a similar extent as TAPBPR^WT (Figure 2a). This may be due, in part, to TAPBPR enhancing the trafficking of associated, peptide-receptive MHC I with it through the secretory pathway. However, when we tested the ability of TAPBPR to mediate peptide dissociation (i.e. removal of the bound fluorescent peptide in the presence of an unlabelled competitor peptide) (Figure 2b), in contrast to the efficient exchange observed with TAPBPR^WT, very little, if any, dissociation of fluorescent peptide was observed with TAPBPR^Øloop (Figure 2c–2e). This suggests that the K22-D35 loop of TAPBPR is essential for mediating efficient peptide dissociation from HLA-A*68:02.

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Next, we tested the ability of both plasma membrane bound TAPBPR^L30G and TAPBPR^ØG30L to promote peptide exchange in our assay. As observed above with TAPBPR^Øloop, plasma membrane expressed TAPBPR^L30G and TAPBPR^ØG30L were able to promote the association of exogenous fluorescent peptide onto cells (Figure 2a). However, when we explored the ability of TAPBPR^L30G to mediate fluorescent peptide dissociation from HLA-A*68:02, we found that it was incapable of mediating efficient peptide dissociation in the presence of unlabelled competitor (Figure 2c–2e). Strikingly, alteration of this single residue had the same effect on the function of TAPBPR as mutating the entire loop. The crucial role of L30 in mediating peptide dissociation was further supported with our observation that TAPBPR^ØG30L, in which the leucine residue alone is restored into the dysfunctional TAPBPR^Øloop molecule, transformed it into a functioning peptide exchange catalyst on HLA-A*68:02, albeit at reduced capability compared to TAPBPR^WT (Figure 2c–2e). While TAPBPR^WT efficiently promoted rapid peptide exchange at very low concentrations of competitor peptide, both TAPBPR^Øloop and TAPBPR^L30G were extremely inefficient at mediating peptide exchange on HLA-A*68:02 molecules, even in the presence of high concentrations of competitor peptide (Figure 2e). These results suggest that L30 is a critical residue within the loop of TAPBPR for mediating peptide dissociation from HLA-A*68:02.

We have also recently shown that soluble TAPBPR, consisting only of the lumenal domains of the wild-type molecule (i.e. lacking its transmembrane region and cytoplasmic tail) can also efficiently promote peptide exchange on three MHC I molecules: HLA-A*68:02, HLA-A2 and H-2K^b (Ilca et al., 2018). Therefore, we can also use soluble TAPBPR as a second means to assay the ability of TAPBPR mutants to mediate efficient peptide exchange on MHC I. Using this approach, the majority of MHC I molecules that TAPBPR will have access to will be folded with bound peptides of relativity high affinity expressed on the surface of cells. The lumenal domains of the TAPBPR variants with C-terminal His-tags were purified using Ni-affinity chromatography from the culture supernatants of transfected 293T cells (Figure 3a). Differential scanning fluorimetry revealed all TAPBPR loop variants had a similar melting temperature as TAPBPR^WT (Figure 3b), indicating the alterations made to the loop had not significantly affected protein folding and stability. Comparison of the ability of the soluble TAPBPR variants to mediate peptide exchange on HLA-A*68:02 molecules on HeLaM cells revealed that TAPBPR^WT was most efficient, followed by TAPBPR^ØG30L, which exhibited ~33% peptide exchange activity relative to TAPBPR^WT (in the presence of 10 nM ETVSK*QSNV) (Figure 3c and d). However, both TAPBPR^L30G and TAPBPR^Øloop were unable to efficiently mediating peptide exchange, displaying only ~3% of the exchange activity of TAPBPR^WT (in the presence of 10 nM ETVSK*QSNV) (Figure 3c and d). As previously shown (Ilca et al., 2018), soluble TAPBPR^TN5, a mutant which cannot bind to MHC I, did not mediate any peptide exchange (Figure 3c and d). This hierarchy (WT>ØG30L>L30G>TN5) was maintained over a wide range of exogenous TAPBPR concentrations (Figure 3e). Furthermore, the same hierarchical order of peptide exchange efficiency for the variants was observed using another HLA-A*68:02-binding peptide, YVVPFVAK*V (Figure 3c & d).

Figure 3

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When we determined the ability of the soluble TAPBPR variants to bind to HLA-A*68:02, TAPBPR^WT was the most efficient binder, followed by TAPBPR^ØG30L (Figure 4a). However, TAPBPR^L30G and TAPBPR^Øloop were unable to make productive interactions with surface expressed HLA-A*68:02 (Figure 4a). As expected, TAPBPR^TN5, which has a disrupted binding site for MHC I, was unable to bind to cells (Hermann et al., 2013) (Figure 4a). As this inability of soluble TAPBPR^Øloop and TAPBPR^L30G to interact with MHC I contradicted our finding with their membrane-bound counterparts (Figure 1e), we determined whether these soluble TAPBPR variants could bind to the total cellular MHC I from cell lysates in pull-down experiments. All soluble TAPBPR variants (with the exception of TAPBPR^TN5) were capable of binding significant amounts of MHC I, although less MHC I was detected bound to TAPBPR^L30G and TAPBPR^Øloop (Figure 4b). A major difference between surface MHC I on intact cells (Figure 4a) and total cellular MHC I in detergent (Figure 4b) is the availability of peptide-receptive molecules. Therefore, we speculated that TAPBPR molecules with alterations to the loop are still able to bind to peptide-receptive MHC I but are unable to physically make MHC I peptide-deficient due to their inability to dissociate peptide. Consistent with this, incubation of cells at 26°C, which increases the expression of peptide-receptive MHC I on the plasma membrane (Ljunggren et al., 1990; Schumacher et al., 1990; Garstka et al., 2015), resulted in the TAPBPR loop variants, but not TAPBPR^WT, exhibiting increased binding to cells (Compare Figure 4c with Figure 4a summarised in Figure 4d). While no significant change was observed in the ability of exogenous TAPBPR^WT to bind to cells incubated at the lower temperature, a ~ 7 fold increase in binding of TAPBPR^L30G and TAPBPR^Øloop was observed at 26°C compared to 37°C, and the binding of TAPBPR^ØG30L increased by ~2.5 fold (Figure 4d). When given access to surface expressed peptide-receptive MHC I upon incubation at 26°C, all soluble TAPBPR variants exhibited a corresponding increase in the ability to mediate peptide exchange (compare Figure 3c (37°C) with Figure 4e (26°C), summarised in Figure 4f). These findings suggest that TAPBPR variants lacking L30 are indeed able to bind to peptide-receptive MHC I but are unable to physically make MHC I peptide-deficient due to their inability to efficiently facilitate peptide dissociation. Consistent with this, when cells cultured at 26°C were incubated with an HLA-A*68:02-binding peptide prior to testing soluble TAPBPR binding, the peptide significantly reduced the ability of TAPBPR^Øloop to bind to cells, while the binding of TAPBPR^WT was unaffected (Figure 4—figure supplement 1).

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Having shown a role for the K22-D35 loop of TAPBPR in mediating efficient peptide dissociation from MHC I, we next determined whether the peptide repertoire presented by MHC I molecules was altered in cells upon mutation of the loop. When we compared the immunopeptidomes of IFNγ-treated HeLaM-TAPBPR^KO cells expressing TAPBPR^WT with cells expressing TAPBPR^Øloop, we found significant changes in the peptides presented on MHC I (Figure 5a). 461 peptides were found exclusively in the TAPBPR^WT-expressing cells and 550 peptides were found exclusively in the cells expressing the Øloop variant (Figure 5a). Label-free quantitation by mass spectrometry revealed that there were also significant changes in the abundance of some peptides between TAPBPR^WT and TAPBPR^Øloop, with 193 peptides exhibiting increased abundance in cells expressing TAPBPR^WT (Red circles, Figure 5b) and 222 peptides displaying increased abundance in TAPBPR^Øloop-expressing cells (Blue circles, Figure 5b). These findings demonstrate that there are significant changes in the peptide repertoire presented on MHC I upon mutation of the TAPBPR loop. While there were still large differences in the peptide repertoires between TAPBPR^WT and TAPBPR^ØG30L-expressing cells, based on a presence/absence approach (Figure 5a), restoration of the leucine residue into the loop appeared to somewhat reduce some of the changes observed in peptide abundance (Figure 5b). Assignment of the identified peptides to the MHC I allotype found in HeLaM cells revealed a similar HLA distribution between the various loop mutants (Figure 5c and d).

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Figure 5—source data 1

Binding of TAPBPR to the individual HLA molecules found in HeLaM cells.

Soluble TAPBPR^WT or soluble TAPBPR^Øloop were incubated with the indicated LABScreen single antigen HLA class I beads (from One Lambda, Canoga Park, California) for 60 min at RT. After washing 3 times in PBS, bound TAPBPR was detected using the TAPBPR-specific mAb PeTe4 and a goat anti-mouse PE-conjugated secondary antibody. Samples were analysed using the Luminex Fluoroanalyser system. The table shows the MFI of PeTe4 staining ± SD for the indicated HLA beads incubated in the absence and presence of 1 µM of TAPBPR^WT or TAPBPR^Øloop. The expression of MHC I (detected using W6/32) provided by One Lambda for the specific lot of LABScreen beads used is included as a control.

https://doi.org/10.7554/eLife.40126.014

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Figure 5—source data 2

Dataset 1 - peptides eluted from W6/32-reactive MHC I complexes from IFNγ treated HeLaM-TAPBPR^KO cells expressing TAPBPR^WT.

https://doi.org/10.7554/eLife.40126.015

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Figure 5—source data 3

Dataset 1 - peptides eluted from W6/32-reactive MHC I complexes from IFNγ treated HeLaM-TAPBPR^KO cells expressing TAPBPR^Øloop.

https://doi.org/10.7554/eLife.40126.016

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Figure 5—source data 4

Dataset 1 - peptides eluted from W6/32-reactive MHC I complexes from IFNγ treated HeLaM-TAPBPR^KO cells expressing TAPBPR^ØG30L.

https://doi.org/10.7554/eLife.40126.017

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Figure 5—source data 5

Dataset 2 - peptides eluted from W6/32-reactive MHC I complexes from IFNγ treated HeLaM-TAPBPR^KO cells expressing TAPBPR^WT.

https://doi.org/10.7554/eLife.40126.018

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Figure 5—source data 6

Dataset 2 - peptides eluted frm W6/32-reactive MHC I complexes from IFNγ treated HeLaM-TAPBPR^KO cells expressing TAPBPR^Øloop.

https://doi.org/10.7554/eLife.40126.019

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Figure 5—source data 7

Dataset 2 - peptides eluted from W6/32-reactive MHC I complexes from IFNγ treated HeLaM-TAPBPR^KO cells expressing TAPBPR^ØG30L.

https://doi.org/10.7554/eLife.40126.020

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Figure 5—source data 8

Dataset 1 - analysis of eluted peptides used to generate volcano plots.

https://doi.org/10.7554/eLife.40126.021

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Figure 5—source data 9

Dataset 2 - analysis of eluted peptides used to generate volcano plots.

https://doi.org/10.7554/eLife.40126.022

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Figure 5—source data 10

Peptides eluted from W6/32-reactive MHC I complexes from IFNγ treated HeLaM-TAPBPR^KO cells expressing TAPBPR^M29.

https://doi.org/10.7554/eLife.40126.023

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Figure 5—source data 11

Dataset 3 - peptide list for third biological repeat for TAPBPR^WT expressing cells.

https://doi.org/10.7554/eLife.40126.024

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Figure 5—source data 12

Dataset 3 - peptide list for third biological repeat for TAPBPR^Øloop expressing cells.

https://doi.org/10.7554/eLife.40126.025

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Figure 5—source data 13

Dataset 3 - peptides list for third biological repeat for TAPBPR^ØG30L expressing cells.

https://doi.org/10.7554/eLife.40126.026

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Given the ability of surface expressed TAPBPR^WT, but not of TAPBPR^Øloop, to mediate peptide dissociation from surface MHC I molecules, as identified in Figure 2, we performed an additional experiment in which the cells were incubated at 37°C in media after harvesting to permit the surface expressed TAPBPR (~5% of the total TAPBPR pool in these cells [Ilca et al., 2018]) to perform peptide dissociation/exchange on surface expressed MHC I molecules, prior to performing immunopeptidomic analysis. While this experiment confirmed the significant changes in the peptide repertoire presented on MHC I upon mutation of the TAPBPR loop (Figure 5e and f), it also surprisingly revealed a large effect on the peptides assignable to HLA-A*68:02 between cells expressing TAPBPR^WT and TAPBPR^Øloop (Figure 5g and h). Based on a presence/absence approach, only 29% in TAPBPR^WT were now assignable to HLA-A*68:02, compared to 37% for cell expressing TAPBPR^Øloop-expressing cells (which was similar to the results found when the immunopeptidomics was performed immediately after cell harvesting [Figure 5d]). Similarly, abundance analysis revealed >80% of the up-modulated peptides in TAPBPR^Øloop-expressing cells belonged to HLA-A*68:02, compared to only ~20% of the up-modulated peptides in TAPBPR^WT (Figure 5h). In keeping with this, we observed similar alterations in HLA-A*68:02-assignable peptides when the leucine residue was restored into the dysfunctional loop (Figure 5g and h), as the ones observed for TAPBPR^WT. This suggests that TAPBPR with a functional loop is preferentially stripping a proportion of peptides from HLA-A*68:02 molecules, but not from the HLA-B or -C molecules found on HeLaM cells. To explore this further, we compared the ability of soluble TAPBPR to bind to soluble heterotrimeric HLA-A*68:02, -B*15:03 or -C*12:03 molecules coupled to beads. This revealed a strong interaction between TAPBPR^WT and HLA-A*68:02 (Figure 5—source data 1). However, soluble TAPBPR^WT failed to bind to heterotrimeric HLA-B*15:03 and –C*12:03 molecules (Figure 5—source data 1). This preferential association of TAPBPR with peptide-loaded HLA-A*68:02 over the other MHC I found in HeLaM cells likely explains why we only observe a loss of HLA-A*68:02 peptides in cells expressing TAPBPR with a functional loop. Interestingly, this analysis of soluble HLA molecules coupled on beads also confirmed the considerable reduction in binding of TAPBPR to HLA-A*68:02 upon mutation of the loop (Figure 5—source data 1). Taken together, this data demonstrates that the loop is involved in selecting MHC I peptides within cells and highlights the importance of leucine 30 in the peptide selection process, particularly for HLA-A*68:02 molecules.

Given the proximity of the TAPBPR loop to the F pocket of MHC I (Figure 1) and that the L30 residue of TAPBPR was both necessary and sufficient for efficient peptide exchange on HLA-A*68:02 (Figure 2 and 3), we hypothesised that the TAPBPR loop facilitates peptide dissociation by binding into the F pocket, thereby competing with the C-terminus of the peptide. If so, this competitive binding would only be possible for MHC I molecules that could accommodate leucine or similar hydrophobic residues in the F pocket. With this in mind, we explored the importance of the TAPBPR K22-D35 loop in mediating peptide exchange on two other MHC I molecules, HLA-A*02:01 and H-2K^b, which naturally accommodate hydrophobic amino acids in their F pocket like HLA-A*68:02. Compared to HLA-A*68:02, HLA-A*02:01 accommodates very similar anchor residues in both B and F pockets, whereas H-2K^b has a completely different binding motif, with the exception that it binds a similar anchor residue in the F pocket (Figure 6a). Thus, given that both HLA-A*02:01 and H-2K^b could potentially accommodate L30 of the TAPBPR loop in their F-pocket, we predicted TAPBPR-mediated peptide editing on these two additional MHC I would also be dependent on L30.

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As observed above with HLA-A*68:02, we found that TAPBPR^WT was the most efficient catalyst on both HLA-A*02:01 and H-2K^b, followed by TAPBPR^ØG30L, while both TAPBPR^L30G and TAPBPR^Øloop were least efficient (Figure 6b and c). For example, the Øloop, L30G, and ØG30L variants exhibited ~23%, 32% and 54% activity relative to TAPBPR^WT, respectively, when measuring NLVPK*VATV binding onto HLA-A2 introduced into HeLaM-HLA-ABC^KO cells by transduction (Figure 6c). We observed similar trends using another HLA-A2 binding peptide, YLLEK*LWRL, on this cell line, as well as when testing peptide binding to HLA-A2 molecules expressed on MCF-7 cells (Figure 6—figure supplement 1). For SIINKEK*L binding to H-2K^b expressed on EL4 cells, the Øloop, L30G, and ØG30L variants exhibited ~30%, 31%, 85% activity relative to TAPBPR^WT, respectively (Figure 6c). Thus, although TAPBPR can still mediate peptide exchange on HLA-A2 and H-2K^b in the absence of the loop to some extent, the L30 residue in the K22-D35 loop of human TAPBPR plays a critical role in promoting efficient peptide exchange on HLA-A*68:02, HLA-A2 and H-2K^b. The shared dependency on L30 to enable TAPBPR to efficiently mediate peptide exchange on both H-2K^b and HLA-A*68:02 is remarkable, considering the low degree of similarity between these two MHC I, both in their amino acid sequences and in their binding motifs. However, the key shared feature between these MHC I is that both accommodate hydrophobic residues in their F pocket.

We next tested the role of the TAPBPR loop in catalysing peptide exchange on an MHC I molecule that does not accommodate a hydrophobic residue in its F pocket. In order to only explore the contributing role of the F pocket, with minimal other polymorphisms, we chose HLA-A*68:01, since it only differs from HLA-A*68:02 by five amino acids (Niu et al., 2013). Most of the differences between these two HLA are residues dictating the specificity of the F pocket for the C-terminal peptide anchor residue. Hence, while both molecules have a similar anchor residue in their B pocket, the F pocket of HLA-A*68:02 accommodates an aliphatic residue, whereas the F pocket of HLA-A*68:01 accommodates a basic residue (Figure 6a) (Niu et al., 2013). Strikingly, in contrast to the three MHC I molecules tested previously (which all bind hydrophobic residues in their F pocket), there was a significant impairment in the ability of TAPBPR^WT to load peptides onto HLA-A*68:01 (Figure 6b and c). Moreover, there was no significant difference in the ability of TAPBPR to exchange peptides on HLA-A*68:01 upon mutation of the loop, with the different loop variants all exhibiting a similar ability as soluble TAPBPR^WT to load the fluorescent peptide KTGGPIYK*R onto HLA-A*68:01 expressed on HeLaM-HLA-ABC^KO cells (Figure 6b and c). Thus, in contrast to HLA-A*68:02, which is extremely receptive to TAPBPR-mediated peptide exchange in a loop-dependent manner, HLA-A*68:01 is significantly less responsive to TAPBPR-mediated peptide exchange, which only occurs in a loop-independent manner. Therefore, the five amino acid differences between these two HLA molecules, three of which are around the F pocket (Figure 6d), strongly influence the receptivity of these two MHC I molecules to TAPBPR-mediated peptide exchange, the efficiency of which appears to be strongly influenced by the ability of the K22-D35 loop (specifically L30) to interact with the peptide binding groove. These findings support the concept that the L30 residue of TAPBPR is capable of binding into the F pocket of MHC I in order promote the dissociation of the bound peptide in a competitive manner.

Our data thus far is consistent with the L30 residue of the TAPBPR loop binding to the F pocket of MHC I molecules which accommodate hydrophobic residues. One residue that differs between HLA-A*68:02 and -A*68:01 and was reported to be crucial in determining the F pocket specificity of MHC I for peptide residues is the one on position 116 (Sidney et al., 2008). The impact of residue 116 on the F pocket architecture is well highlighted in the crystal structures of peptide-bound HLA-A*68:02 and HLA-A*68:01 (Niu et al., 2013). Namely, HLA-A*68:02 contains a tyrosine on position 116, whereas HLA-A*68:01 has an aspartate (Figure 7a). As captured in the two structures, D116 of HLA-A*68:01 forms strong dipole interactions with both residue R114 of the groove and with the arginine on position 9 of the peptide, determining a strong preference of the HLA-A*68:01 F pocket for basic anchor residues. In contrast, residue Y116 of HLA-A*68:02 does not form any interaction with H114, keeping the hydrophobic patches of the F pocket in an open conformation, which allows it to accommodate hydrophobic peptide residues (Figure 7a). We believe that this is the reason why the L30 residue of the TAPBPR loop has access to the F pocket of HLA-A*68:02, but not to the one of HLA-A*68:01.

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To explore whether the F pocket specificity of MHC I molecules was indeed crucial for allowing TAPBPR-mediated peptide exchange, we altered the F pocket of both HLA-A*68:02 and -A*68:01, by switching their residues on position 116, producing HLA-A*68:02^Y116D and HLA-A*68:01^D116Y. We subsequently tested their ability to bind soluble TAPBPR. When transduced into HeLaM-HLA-ABC^KO cells, the 116-mutated HLA-A68 molecules were expressed at equivalent levels as their WT counterparts (Figure 7b). When we tested the ability of soluble TAPBPR^WT to bind to cells expressing the HLA-A*68:02 molecules, strikingly, TAPBPR exhibited extremely low levels of binding to HLA-A*68:02^Y116D compared to HLA-A*68:02^WT molecules (Figure 7c and d). Conversely, TAPBPR binding to HLA-A*68:01 was significantly enhanced upon mutation of residue 116 from D to Y (Figure 7c and d). The influence of altering the F pocket on TAPBPR binding was further verified on another MHC I molecule, HLA-A2 (Figure 7—figure supplement 1). Together, these results demonstrate that the binding of TAPBPR to MHC I molecules is directly influenced by the architecture of the F pocket.

Next, we tested the ability of TAPBPR^WT to mediate peptide exchange on the same panel of HLA-A68 molecules. Since HLA-A*68:01 and -A*68:02 have similar residues on position 114, we hypothesized that swapping their 116 residues might also swap their F pocket specificities. Thus, we interrogated the ability of these molecules to bind both the HLA-A*68:02-specific peptide ETVSK*QSNV, as well as the HLA-A*68:01-specific peptide KTGGPIYK*R, in the presence or absence of soluble TAPBPR. While TAPBPR^WT efficiently mediated the loading of ETVSK*QSNV onto HLA-A*68:02^WT molecules, it failed to load this peptide onto HLA-A*68:02^Y116D molecules (Figure 7e and f). Similarly, TAPBPR^WT failed to load ETVSK*QSNV onto HLA-A*68:01^WT molecules, however, efficiently loaded this peptide onto HLA-A*68:01^D116Y (Figure 7e and f). Moreover, we found that while TAPBPR^WT could not load KTGGPIYK*R onto HLA-A*68:02^WT molecules, it could efficiently load this peptide onto HLA-A*68:02^Y116D (Figure 7g and h). Likewise, while HLA-A*68:01^WT could bind KTGGPIYK*R in the presence of TAPBPR^WT, no loading of this peptide was observed onto HLA-A*68:01^D116Y (Figure 7g and h). These results are in keeping with the fact that the alterations made to the F pocket of the HLA-A68 molecules, and more specifically, to residue 116 alone, have altered peptide specificity. Nonetheless, it is interesting that although low levels of TAPBPR binding are observed to both HLA-A*68:02^Y116D and HLA-A*68:01^WT, TAPBPR is still capable of mediating peptide exchange on these molecule (albeit to a significantly lower extent as compared to their Y116-containing counterparts) and of loading the correct peptide based on their F pocket anchor.

Dockings of TAPBPR with MHC I were carried out prior to the recent structure determination of the TAPBPR:MHC I complex structures. For the docking studies, we used our previously determined model for human TAPBPR based on the structure of tapasin (Hermann et al., 2013) and human HLA-A2 (Saper et al., 1991) (PDB ID 3HLA). TAPBPR was manually docked onto MHC I with the following restraints: (i) the membrane-proximal domains of each component, α3 of MHC I and the IgC domain of TAPBPR were oriented so as to allow these proteins to maintain complex formation when membrane-embedded; (ii) the surface shape complementarity of the components was maintained; (iii) the side chain of residue T134 of MHC I was kept within 10 Å of the sidechains of residues E205-Q209 (TN6 patch) and I261 (TN5 patch) on the IgV domain of TAPBPR; and (iv) sidechains of E222 and D223 of MHC I were kept within 10 Å of the sidechains of residues R335 (TC2) and Q336-S337 (TC3) on the IgC domain of TAPBPR. These last two restraints being mutations known to knock out the interaction in cells (Hermann et al., 2013) and the relatively loose restraints allowing for some conformational flexibility of the components upon complex formation.

HeLaM cells, a variant HeLa cell line that is more responsive to IFN (Tiwari et al., 1987) (a gift from Paul Lehner, University of Cambridge, UK), their modified variants, and HEK-293T (from Paul Lehner, University of Cambridge, UK) were maintained in Dulbecco's Modified Eagle's medium (DMEM; Sigma-Aldrich, UK), supplemented with 10% fetal calf serum (Gibco, Thermo Fisher Scientific), 100 U/ml penicillin and 100 µg/ml streptomycin (Gibco, Thermo Fisher Scientific) at 37°C with 5% CO₂. All cells were confirmed to be mycoplasma negative (MycoAlert, Lonza, UK). Authenticity of HeLaM was verified by tissue typing for HLA molecules and by the continuous confirmation that these cell lines had the expected HLA class I tissue type monitored by staining with specific HLA antibodies and by mass spectrometry.

TAPBPR^WT and TAPBPR^TN5 constructs cloned in the lentiviral vector pHRSIN-C56W-UbEM have previously been described (Boyle et al., 2013; Hermann et al., 2013). TAPBPR^Øloop, in which amino acids comprising the 22–35 loop were replaced with amino acids glycine, alanine and serine, was created from TAPBPR^WTpHRSIN-C56W-UbEM using the following procedure: First, amino acids 22–28 were replaced using quick-change site-directed mutagenesis using primers M22-for and M22-rev (see Table 2 for primer sequences). Subsequently, amino acids 29–35 were replaced using a two-step PCR. For this, the TAPBPR insert was amplified in two separate pieces, starting from each side of the mutation site (primers TAPBPR^WT-BamHI-for and M29-rev for the N terminus-containing side and primers M29-for and TAPBPR^WT-NotI-rev for the C terminus-containing side). Subsequently, the two pieces bearing complementary regions over the mutated site were used in a second PCR reaction to amplify the whole TAPBPR mutated insert using primers TAPBPR^WT-BamHI-for and TAPBPR^WT-NotI-rev. TAPBPR^L30G and TAPBPR^ØG30L were generated from TAPBPR^WT and TAPBPR^Øloop respectively, using primers L30G-for and L30G-rev or ØG30L-for and ØG30L-rev, by quick change site-directed mutagenesis.

The HLA-A*68:02^WT construct was cloned by consecutive rounds of quick-change site-directed mutagenesis, using the HLA-A*68:01^WT construct as a template (see Table 3 for primer sequences). Since residue 116 was mutated last in this process, the HLA-A*68:02^Y116D mutant was the final intermediate in this cloning process. The HLA-A*68:01^D116Y was cloned by quick-change site-directed mutagenesis, using primers A6801-D116Yonly-Fwd and A6801-D116Yonly-Rev. The HLA-A2^Y116D mutant was cloned by quick-change site-directed mutagenesis, using primers A2-Y116D-Fwd and A2-Y116D-Rev.

Reconstitution of the TAPBPR variants into the TAPBPR-knockout HeLaM cell line (HeLaM-TAPBPR^KO), and HLA into the HeLaM-HLA-ABC^KO cells was performed using lentiviral transduction and the cells were subsequently cultured as previously described (Neerincx et al., 2017; Neerincx and Boyle, 2018). Cells were induced with 200 U/ml IFN-γ (Peprotech, UK) for 48–72 hr where indicated.

To make secreted forms of the TAPBPR loop variants enumerated above, the lumenal domains were cloned into a modified version of the PB-T-PAF vector where the N-terminal Protein A fusion was removed and a C-terminal His₆ tag introduced and expressed in 293 T cells using the PiggyBac expression system (Li et al., 2013). 48 hr after transfection, cells were transferred for at least 5 days into selection media (DMEM supplemented with 10% FBS, 1% pen/strep, 3 µg/mL puromycin (Invivogen, San Diego, CA) and 700 µg/mL geneticin (Thermo Fisher Scientific, UK). To induce protein expression, cells were harvested and transferred into DMEM supplemented with 5% FBS, 1% pen/strep and 2 µg/mL doxycycline (Sigma-Aldrich, UK). After 5–7 days, the media was collected and TAPBPR was purified using Ni-NTA affinity chromatography. For purity assessment, elution fractions were analysed by SDS-PAGE, followed by Coomassie staining

Thermofluor experiments were performed in 96-well low profile clear PCR plates for Viia7 cyclers (Axygen). Reactions of 20 μl comprised of 5 μg protein, 1x protein Thermal Shift Dye (Life Technologies) in PBS pH 7.4. The melting curve was performed using a Viia7 thermocycler between 20°C and 95°C in 1°C steps with 20 s equilibration time per step and fluorescence monitored on the ROX channel.

HLA class I molecules were isolated from HeLaM-TAPBPR^KO cells transduced with either TAPBPR^WT, TAPBPR^Øloop or TAPBPR^ØG30L using standard immunoaffinity chromatography employing the pan-HLA class I-specific antibody W6/32 (produced in-house), as described previously (Kowalewski and Stevanović, 2013). Tissue typing confirmed the HeLaM cells express HLA-A*68:02, -B*15:03 and –C*12:03.

Isolated HLA peptides were analysed in five technical replicates. Peptide samples were separated by nanoflow high-performance liquid chromatography (RSLCnano, Thermo Fisher Scientific, Waltham, MA) using a 50 μm × 25 cm PepMap rapid separation liquid chromatography column (Thermo Fisher Scientific) and a gradient ranging from 2.4% to 32.0% acetonitrile over the course of 90 min. Eluting peptides were analyzed in an online-coupled LTQ Orbitrap XL mass spectrometer (Thermo Fisher Scientific) using a top five CID (collision-induced dissociation) fragmentation method.

Spectra were annotated to corresponding peptide sequences by database search of the human proteome as comprised in the Swiss-Prot database (20,279 reviewed protein sequences, September 27^th 2013) by employing the SEQUEST HT search engine (Eng et al., 1994) (University of Washington) integrated into ProteomeDiscoverer 1.4 (Thermo Fisher Scientific). Data processing was performed without enzyme specificity, with peptide length limited to 8–12 amino acids, and methionine oxidation set as dynamic modification. The false discovery rate was calculated by the Percolator algorithm (Käll et al., 2007) and set to 5%. HLA annotation was performed using NetMHCpan-4.0 with a percentile rank threshold of 2%.

We used label-free quantitation (LFQ) as described previously (Nelde et al., 2018) to assess the relative HLA ligand abundances between TAPBPR^WT, TAPBPR^Øloop or TAPBPR^ØG30L expressing cells. Briefly, relative quantitation of HLA ligands was performed by calculating the area under the curve of the respective precursor extracted ion chromatogram (XIC) using ProteomeDiscoverer 1.4 (Thermo Fisher Scientific). For LFQ analysis the total injected peptide amount of all samples was normalised prior to LC-MS/MS analysis. Volcano plots were computed using an in-house R script (v3.2) and depict pairwise comparisons of the ratios of the mean areas for each individual peptide in the five LFQ-MS runs. Significant modulation was defined by an adjusted p-value of < 0.01 and a fold change of ≥ log₂ 2 fold change, as calculated by two-tailed t-tests implementing Benjamini-Hochberg correction.

The following fluorescent MHC I-specific peptides were used (K* represents a lysine labelled with 5-carboxytetramethylrhodaime [TAMRA]): ETVSK*QSNV (HLA-A*68:02), YVVPFVAK*V (HLA-A*68:02), NLVPK*VATV (HLA-A*02:01), YLLEK*LWRL (HLA-A*02:01), SIINFEK*L (H-2K^b), and KTGGPIYK*R (HLA-A*68:01). The following unlabeled competitor peptides were used: YVVPFVAKV, which exhibits high affinity of HLA-A*68:02 and EGVSEQSNG, a non-binding derived of ETVSEQSNV, obtained by replacing its anchor residues (amino acids on positions 2 and 9) with glycine. All peptides were purchased from Peptide Synthetics, UK.

TAPBPR was detected using either PeTe4, a mouse monoclonal antibody (mAb) specific for the native conformation of TAPBPR, raised against amino acids 22–406 of human TAPBPR (Boyle et al., 2013) that does not cross-react with tapasin (Hermann et al., 2013), or ab57411, a mouse mAb raised against amino acids 23–122 of TAPBPR that is reactive to denatured TAPBPR (Abcam, UK). UGT1 was detected using the rabbit mAb ab124879 (Abcam). MHC class I heavy chains were detected using mAb HC10 (Stam et al., 1986). Soluble TAPBPR variants were detected using the mouse anti-polyhistidine mAb H1029 (Sigma-Aldrich). A mouse IgG2a isotype control was also used as a control (Sigma-Aldrich).

Following trypsinisation, cells were washed in 1% bovine serum albumin (BSA), dissolved in 1x phosphate-buffered saline (PBS) at 4°C, and then stained for 30 min at 4°C in 1% BSA containing with PeTe4 or with an isotype control antibody. After washing the cells to remove excess unbound antibody, the primary antibodies bound to the cells were detected by incubation at 4°C for 25 min with either goat anti-mouse Alexa-Fluor 647 IgG (Invitrogen Molecular Probes, Thermo Fisher Scientific). After subsequent three rounds of washing, the fluorescence levels were detected using a BD FACScan analyser with Cytek modifications and analysed using FlowJo (FlowJo, LLC, Ashland, OR).

For peptide binding in the presence of recombinant TAPBPR, the cells were treated with or without the indicated concentration of recombinant TAPBPR (unless otherwise indicated, we used 100 nM for HLA-A*68:02, 1 µM used for HLA-A*02:01, H-2K^b and HLA-A*68:01). After 15 min, the desired TAMRA-labelled peptide was added to the cells and incubated at 37°C (15 min for HLA-A*68:02, 60 min for HLA-A*02:01 and –A*68:01, or 30 min for H-2K^b). For experiments performed at 26°C, following cell seeding and IFNγ stimulation at 37°C for 48 hr, cells were transferred at 26°C for another 12 hr to allow for the expression of sub-optimally loaded MHC I molecules at the cell surface. The TAPBPR and peptide binding was then performed as described above, with all incubation steps being performed at 26°C instead of 37°C (with the exception of cell trypinisation which was carried out at 37°C). In cases where the peptide binding was facilitated by over-expressed TAPBPR, the labelled peptide was directly added to the cells, without using recombinant TAPBPR. Following the peptide treatment, the cells were washed three times in 1x PBS and harvested. The level of bound peptide/cell was determined by flow cytometry, using the YelFL1 channel (Cytek).

IFNγ-treated HeLaM-TAPBPR^KO cell lines reconstituted with the different TAPBPR variants, were treated with 10 nM TAMRA-labelled peptide of interest diluted in opti-MEM for 15 min at 37°C, as described above. Following the binding step, the peptide-containing media was removed, the cells were washed and then treated with media alone or with different concentrations of non-labelled peptide for another 15 min at 37°C. The cells were then washed and harvested and the level of bound peptide per cell was determined by flow cytometry, using the YelFL1 channel (Cytek).

For TAPBPR immunoprecipitation experiments from cells over-expressing the panel of TAPBPR variants, cells were lysed in 1% triton X-100 (VWR, Radnor, PN), Tris-buffered saline (TBS) (20 mM Tris-HCl, 150 mM NaCl, 2.5 mM CaCl₂) supplemented with 10 mM N-ethylmaleimide (NEM), 1 mM phenylmethylsulfonyl fluoride (PMSF) (Sigma-Aldrich), and protease inhibitor cocktail (Roche, UK) for 30 min at 4°C. Nuclei and cell debris were pelleted by centrifugation at 13,000 × g for 15 min and supernatants were collected. Immunoprecipitation was performed with the TAPBPR-specific mAb PeTe4 (Boyle et al., 2013) coupled to protein A sepharose (GE Healthcare), at 5 µg antibody per sample, for 2 hr at 4°C with rotation. Following immunoprecipitation, beads were washed thoroughly in 0.1% detergent-TBS to remove.

For pulldown experiments using soluble TAPBPR proteins, IFNγ-stimulated HeLa-TAPBPR^KO cells were harvested, lysed and cleared of cell debris as above. In order to remove cellular factors which bind non-specifically to the sepharose beads, the cell lysate was pre-cleared by treatment with protein A sepharose alone, for 30 min at 4°C. Subsequently, the lysate was aliquoted and incubated with 5 µg of the soluble TAPBPR variant for 90 min at 4°C. Immunoprecipitation of soluble TAPBPR was performed using PeTe4 as above. Soluble TAPBPR was detected on western blots with the anti-polyHis primary antibody. Gel electrophoresis and western blot analysis was performed as described in Neerincx et al. (2017).

The funders had no role in study design, data collection and interpretation, or the decision to submit the work for publication.

We are extremely grateful to Gemma Brewin and Sarah Peacock (Tissue Typing Laboratory, Cambridge University Hospitals NHS Foundation Trust) for both the use of and their guidance using the LABScreen single antigen HLA class I beads and the Luminex Fluoroanalyser system. We thank Ben Challis and John Trowsdale from the University of Cambridge for proofreading our manuscript and Nico Trautwein from Eberhard Karls University Tübingen for assistance with Immunopeptidomics.

© 2018, Ilca et al.

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