The topology, structure and PE interaction of LITAF underpin a Charcot-Marie-Tooth disease type 1C
- Anita K. Ho†1,
- Jane L. Wagstaff†2,
- Paul T. Manna1,
- Lena Wartosch1,
- Seema Qamar1,
- Elspeth F. Garman3,
- Stefan M. V. Freund2 and
- Rhys C. Roberts1Email author
© Ho et al. 2016
Received: 25 August 2016
Accepted: 16 November 2016
Published: 7 December 2016
Mutations in Lipopolysaccharide-induced tumour necrosis factor-α factor (LITAF) cause the autosomal dominant inherited peripheral neuropathy, Charcot-Marie-Tooth disease type 1C (CMT1C). LITAF encodes a 17 kDa protein containing an N-terminal proline-rich region followed by an evolutionarily-conserved C-terminal ‘LITAF domain’, which contains all reported CMT1C-associated pathogenic mutations.
Here, we report the first structural characterisation of LITAF using biochemical, cell biological, biophysical and NMR spectroscopic approaches. Our structural model demonstrates that LITAF is a monotopic zinc-binding membrane protein that embeds into intracellular membranes via a predicted hydrophobic, in-plane, helical anchor located within the LITAF domain. We show that specific residues within the LITAF domain interact with phosphoethanolamine (PE) head groups, and that the introduction of the V144M CMT1C-associated pathogenic mutation leads to protein aggregation in the presence of PE.
In addition to the structural characterisation of LITAF, these data lead us to propose that an aberrant LITAF-PE interaction on the surface of intracellular membranes contributes to the molecular pathogenesis that underlies this currently incurable disease.
KeywordsLipopolysaccharide-induced tumour necrosis factor-α factor Charcot-Marie-Tooth disease Neuropathy Endosomes
The Charcot-Marie-Tooth diseases (CMT) are among the most common inherited neurological disorders, with an estimated prevalence of 1 in 2500 . CMT is associated with progressive degeneration of peripheral nerves, leading to muscle wasting and weakness, sensory deficits and limb deformities causing significant morbidity across populations .
Peripheral nerves are composed of two main functional anatomical structures: axons and Schwann cells. Axons convey electrical signals to and from the central nervous system and peripheries, while Schwann cells are neural crest-derived cells that engulf peripheral axons providing structural and trophic support. Furthermore, Schwann cells can also be stimulated to wrap multiple layers of plasma membrane around larger axons to form the insulating myelin sheath, a process known as myelination. Consistent with this anatomical and functional dichotomy, CMT can additionally be classified into ‘axonal’ and ‘demyelinating’ forms, reflecting the main sites of cellular dysfunction as the axon or Schwann cell, respectively .
With the advent of next-generation DNA sequencing, mutations in more than 80 genes have now been shown to be associated with CMT, highlighting key genes essential for peripheral nerve function . Of the genes associated with demyelinating CMT, where Schwann cell dysfunction is thought to be the primary underlying pathology, the encoded proteins can be classified into three main groups: structural proteins of the myelin sheath, transcription factors that activate the myelination programme and, interestingly, proteins that are known or predicted to function in intracellular membrane traffic . While Schwann cell-specific expression of CMT-associated proteins explains particular subtypes of demyelinating CMT [6, 7], most CMT-associated genes with known or predicted roles in membrane traffic are widely expressed across diverse tissues and species. Therefore, understanding why mutations in these widely expressed genes lead to isolated demyelinating peripheral neuropathies requires further study at the molecular level.
Here, we present the membrane topology, metal binding and structural features of the human LITAF protein. The biophysical and cell biological effects of the recruitment of interacting proteins to LITAF are then reported, demonstrating the importance of short peptide sequence motifs in the N-terminal proline-rich region for LITAF’s intracellular localisation. Finally, we show that distinct residues contained within the LITAF domain interact with phosphoethanolamine (PE) head groups and that the alteration of these interactions leads to protein aggregation in the presence of a pathogenic CMT1C mutation, providing molecular insights into the underlying pathogenesis of a subtype of this currently incurable, common, inherited disease.
LITAF domains are evolutionarily conserved
LITAF domains are found throughout the eukaryotes, suggesting ancient conserved functions (Additional file 1: Figure S1), with multiple instances of expansion, especially in the metazoa. Of relevance to the current study, one particular duplication appears to have occurred in the common ancestor of the jawed vertebrates (gnathostomes), leading to the eventual existence of two LITAF domain-containing genes in humans: the so-far uncharacterised CDIP1 gene, and the CMT1C-associated gene, LITAF (Additional file 2: Figure S2). Intriguingly, and despite the large lineage-specific expansions seen in the LITAF domain gene family as described, the canonical domain architecture – that is, a proline-rich domain at the N-terminus followed by a C-terminal LITAF domain – is conserved across eukaryotes (with the exception of the apicomplexans), highlighting the likely requirement for both regions to be present for correct intracellular protein function (Additional file 3: Figure S3 and Additional file 4: Figure S4a).
With the aim of identifying key conserved features that are likely to be of critical functional importance, we gathered a cohort of broadly representative LITAF domain sequences from which a comprehensive sequence alignment was derived. This alignment revealed that two pairs of cysteines, located either side of a predicted hydrophobic helix with amphipathic properties, are strictly conserved (Fig. 1 and Additional file 4: Figure S4b, c). These cysteine pairs, in the absence of the intervening helix, are reminiscent of metal-ion coordinating residues found in zinc-finger-like structures . Six additional cysteine residues are present in the predicted hydrophobic helical region of the human LITAF domain, but are less well conserved across our alignment. As their sequence positions are not consistent with the typical known cysteine-containing metal coordinating motifs (e.g. CxxC/HxxC), it appears unlikely that these additional cysteine residues are involved in coordinating a metal ion.
Given that the luminal, transmembrane and cytosolic domains of proteins are conserved to a different extent in evolution [22, 23], the absolute conservation of these residues in tandem led us to hypothesise that the N- and C-termini of the LITAF domain are very unlikely to be separated by a phospholipid bilayer-traversing transmembrane domain as has been previously postulated . Furthermore, most CMT1C-associated pathogenic mutations fall at conserved residues in this pan-eukaryotic alignment of LITAF domains, again highlighting key residues likely to play critical roles in maintaining the function of this ancient domain.
With these points in mind, we set out to re-examine the role of the LITAF domain in targeting the protein to membranes and to experimentally characterise the topology of the human LITAF protein.
LITAF targets to membranes via the hydrophobic helical region
Both the N- and C-termini of LITAF are found on the cytosolic surfaces of vesicles
By analysing the sequences of LITAF domains from Drosophila, Ponting et al.  first postulated that the predicted hydrophobic helical region probably inserted into, but did not traverse, membranes. However, Lee et al.  later assumed that this hydrophobic region within the LITAF domain was a transmembrane helix, thereby concluding that LITAF must be C-terminally inserted into membranes. Importantly, neither hypothesis was supported by experimental data. We therefore used two complementary experimental approaches to determine the membrane topology of LITAF.
Secondly, to verify our findings from the cellular semi-permeabilisation assay, we used a rabbit reticulocyte lysate cell-free system to express HA-LITAF-myc in the presence of microsomes [25, 26]. Once incorporated into microsomal membranes, the susceptibility of both the N- and C-terminal HA and myc epitope tags to protease digestion was ascertained. Using the protection of the processed form of β-lactamase within the microsomal lumen as control, we observed that both the HA and myc tags of LITAF were degraded following incubation with proteinase K (Fig. 3d). These data are therefore consistent with the semi-permeabilisation assay, and are strongly supportive of the hypothesis that both the N- and C- termini of LITAF are found on the cytosolic surface of membranes, and that the LITAF domain is embedded into membranes via the predicted hydrophobic helical region adopting an in-plane membrane anchor configuration .
The LITAF domain coordinates zinc
The LITAF domain consists of cysteine-containing β-hairpins either side of the predicted hydrophobic helical anchor
We next set out to determine the structural characteristics of LITAF. The first NMR experiments of wild type LITAF in an extensive range of membrane-mimicking detergent environments highlighted the crucial importance of the correct choice of solubilising conditions. Under all conditions tested, we observed no signals that we would associate with a structured protein domain. Based on the assumption that these structured residues would be in close association with the membrane, and the high tendency of these samples to form oligomers and aggregate (Additional file 7: Figure S7), we refocused on the LITAF Δ114–139 construct, where the hydrophobic anchor had been removed, thereby rendering this construct soluble in the absence of detergents. 13C/15N-labelled LITAF Δ114–139 protein was straightforwardly expressed and purified from E. coli with an estimated molecular weight of 17 kDa, as determined using a 1D proton transverse relaxation experiment, consistent with a monomeric species in solution.
Unsurprisingly, the LITAF N-helix construct also remained soluble in the absence of detergent. However, similarly to wild type LITAF, the assignment experiments on this construct were also unsuccessful, likely due to the tendency for the N-helix construct to form higher molecular weight oligomers (Additional file 7: Figure S7). Nevertheless, considerable overlap was evident between the N-helix and Δ114–139 constructs in the BEST-TROSY experiments (Additional file 8: Figure S8), suggesting that both LITAF constructs contain very similar secondary structure elements, implying that further interpretation of the Δ114–139 construct would enable key structural motifs of the wild type LITAF protein to be defined.
Building on the near complete assignment of the protein, we proceeded to determine which secondary structural elements are present in the LITAF Δ114–139 construct. This particular approach exploits the exquisite sensitivity of the backbone amino acid chemical shifts to their related secondary structure environment. By using the chemical shift analysis program, TALOS+ , we established that the LITAF domain of the Δ114–139 construct is composed of 5 β-strands: 3 N-terminally to the Δ114–139 deletion site (where the predicted hydrophobic helix would usually reside) and two forming a β-hairpin at the C terminus (Additional file 9: Figure S9a). Supporting these data, the residues assigned to the LITAF domain displayed greater 1H chemical shift distributions (~7.4 to 9.6 ppm), typical of β-sheet residues (Fig. 5), compared to the residues assigned to the N-terminal proline-rich region, which, in contrast, displayed a narrower 1H chemical shift pattern (between 7.8 and 8.4 ppm), suggesting the adoption of a random coil character.
The dynamic qualities of the LITAF Δ114–139 construct were next analysed by performing heteronuclear NOE (hetNOE) experiments. These experiments measure the motion of each assigned residue on a picosecond time scale. Positive hetNOE values close to the value of 1.0 represent slow picosecond motion, which is typically seen when protein domains adopt rigid secondary structure motifs, while small, or even negative, hetNOE values result from fast picosecond motion as seen in unstructured loops and at the flexible termini of proteins. The hetNOE analysis of the Δ114–139 construct, illustrated in (Additional file 9: Figure S9b), demonstrated that four of the five β-strands within the LITAF domain are rigid in the picosecond time scale, while in comparison, the first, most N-terminal β-strand, showed increased motion, and hence flexibility. In contrast, the proline-rich N-terminus showed mainly fast picosecond motion reminiscent of an unstructured protein domain. Interestingly, these analyses also revealed an area of reduced flexibility between residues 22 and 34, a region known to recruit interacting proteins to the LITAF molecule (Additional file 4: Figure S4a) .
NMR-based structural model
The NMR-based model of the LITAF Δ114–139 construct (Fig. 6a), generated from the experimentally determined restraints, illustrate the five β-strands of the LITAF domain: three in an antiparallel orientation at the N-terminus and a β-hairpin at the C-terminus. The conserved cysteine residues, which reside within loops located between β-strands 2 and 3, and between β-strands 4 and 5, are shown to coordinate a zinc atom. Although the five β-strands do not form a continuous sheet, they are in close proximity and are part of an overall compact domain. The glycine/serine linker, that has replaced the predicted hydrophobic in-plane membrane anchor in the LITAF Δ114–139 construct, has been modelled as a short helix by Rosetta. The secondary structure elements are again maintained in the wild type model of the LITAF domain (Fig. 6b), with the predicted hydrophobic in-plane membrane anchor modelled as a single helix by Rosetta. As the coordination of the zinc atom has been maintained, this structure is not as compact as the LITAF Δ114–139 construct, and results in the β-strands lying parallel to the predicted membrane-anchoring helix. The critical coordination of the zinc atom by two pairs of cysteine residues, as shown in both models, is consistent with the rubredoxin, or ‘zinc β-ribbon’ family of zinc finger structural motifs . Efforts to crystallize the LITAF constructs for further structural analysis were unsuccessful.
Recruitment of proteins to the N-terminal proline-rich region directs LITAF to specific endosomal structures
Based on the near complete (89%) assignment of the LITAF Δ114–139 construct, we probed the local interaction of LITAF with the WW domains of NEDD4. A comparison of the BEST-TROSY spectra of the 15N-labelled LITAF Δ114–139 in the presence and absence of recombinantly-expressed NEDD4 596–944 (containing the four WW domains) revealed the loss of a subset of amide cross peaks (Fig. 7b), most often caused by line broadening as a result of micromolar binding affinities. The missing signals correspond to residues 22–29 of the LITAF protein, which contain the PPSY motif known to interact with the WW domains of NEDD4, and are within a region of the N-terminal domain found to be less flexible in the hetNOE experiments. Of note, a second region between residues 57 and 72 of LITAF, which contains a second PPSY motif, showed peak broadening representing a potential additional NEDD4-WW domain interacting site. However, the functional significance of this second PPSY motif is uncertain, since it could not compensate for the loss of the first N-terminal PPSY motif in our pull-down experiments of endogenous NEDD4 from cellular lysates (Fig. 7a). Significantly, we did not detect any consequent structural changes to the C-terminal LITAF domain upon binding of NEDD4 to the N-terminal proline-rich region, showing that these regions of LITAF are structurally independent.
With no apparent structural effect on the C-terminal LITAF domain, we next asked whether the intracellular localisation of LITAF is influenced by the potential recruitment of proteins in vivo. HA-tagged LITAF constructs harbouring mutations that disrupt the PPSY motifs (Y23,61A) were expressed in retinal pigmented epithelial cells (where no endogenously expressed LITAF can be detected) and analysed by immunofluorescence microscopy (Fig. 7c). We found that disruption of the PPSY motifs led to significantly greater colocalisation of LITAF with the plasma membrane marker, wheat germ agglutinin, compared to the wild type protein (Fig. 7d). These data lead us to speculate that, while LITAF domains function as membrane anchors, the specific intracellular localisation is potentially influenced by the recruitment of interacting proteins to the relatively flexible and cytosol-facing N-terminal poly-proline arm.
LITAF interacts specifically with PE head groups
The V144M CMT1C-associated mutation leads to subtle changes in the LITAF Δ114–139 construct and disrupts the interaction with PE
All known CMT1C-associated pathogenic mutations in LITAF are found in the C-terminal LITAF domain. Most mutations reside within the predicted hydrophobic anchor region and are predicted to disrupt the helical propensity of this specific part of the protein (Fig. 1 and Additional file 12: Figure S12a) [40–42]. However, as stated above, experimental study by NMR spectroscopy of constructs containing the predicted hydrophobic helical region were not possible. Nevertheless, the LITAF Δ114–139 construct used to generate the structural model does contain valine at position 144, a residue that, when mutated to a methionine (V144M), is known to cause CMT1C . The V144M mutation does not affect membrane association and its punctate staining pattern by immunofluorescence microscopy is consistent with retained endosomal targeting (Additional file 12: Figure S12b, c). We therefore set out to characterise the effects of this pathogenic mutation by solution NMR spectroscopy. Additional file 11: Figure S11b shows the chemical shift perturbations that are manifested by the V144M mutation in the Δ114–139 construct when compared to the wild type counterpart. If the amino acid substitution had little effect on the construct, we would expect that only signals corresponding to residues in close proximity to the site of mutation would be perturbed. However, dispersed chemical shift perturbations were seen as a result of this CMT1C mutation, including within the region of the N-terminal β-sheets on the opposite side of the deleted hydrophobic anchor. These data therefore suggest that the V144M pathogenic mutation causes subtle biochemical and biophysical changes whilst maintaining the overall secondary structure of the LITAF domain, pointing towards an additional factor that must lead to clinical disease.
By analysing the residues that show the greatest chemical shift perturbations following the introduction of the V144M mutation, we noted that these residues correspond to those that interact with PE head groups (Fig. 8a, b and Additional file 11: Figure S11b). We therefore asked whether the interaction of the LITAF domain with PE is altered by the introduction of the V144M mutation. Indeed, we found that LITAF Δ114–139 V144M interacts with PE at lower head group concentrations (1:250 molar ratio) than the wild type construct whilst utilising the same specific residues (Additional file 13: Figure S13). However, while the V144M construct – similarly to the LITAF Δ114–139 WT protein – remained soluble and un-degraded for periods of many months when stored in isolation, this mutant construct became rapidly insoluble when incubated with PE at 1:1000 molar ratio, leading to the disappearance of the assigned peaks from the NMR spectra and visible precipitation (Fig. 8c and Additional file 13: Figure S13). In contrast, wild type LITAF Δ114–139 remained consistently soluble and folded at these concentrations of PE. To determine the effect of the V144M mutation on the stability of LITAF in vivo, we performed cycloheximide chase experiments in retinal pigment epithelial (RPE) cells stably expressing either wild type HA-LITAF or HA-LITAF V144M. In keeping with our in vitro findings, HA-LITAF harbouring the V144M mutation was degraded more rapidly compared to the wild type construct (Fig. 8d). These data therefore suggest that the V144M pathogenic mutation alters the interaction between the LITAF domain and PE head groups, pointing towards an acquired misfolded state on the surface of membranes, and potentially deleterious effects on intracellular membranes involved in endocytic traffic.
Despite the extensive architectural conservation of LITAF domains across eukaryotes, our report is the first to characterise this ancient domain structurally. In contrast to the assertion made by Lee et al.  that LITAF domains are C-terminally inserted into membranes, our data show that the hydrophobic region does not traverse membranes, demonstrating that both the N-terminal and C-terminal halves of the LITAF domain reside on the cytosolic surface of membranes, in agreement with an earlier proposal made by Ponting et al.  in 2002 and the recent publication by Qin et al. . The LITAF domain consists of five β-sheets, three N-terminal and two C-terminal to the predicted hydrophobic anchor region, and is stabilised by the coordination of a zinc atom by two pairs of evolutionarily-conserved cysteine residues. In contrast, the N-terminal proline-rich region is mostly unstructured and behaves independently of the C-terminal LITAF domain. We show that, far from being an inert component of the full length protein, the ability of the proline-rich region to bind to other cytosolic proteins appears to have a significant influence on intracellular targeting. These data therefore support the notion that, while LITAF domains serve as intracellular membrane anchors, the precise function and localisation is determined by the proteins that interact with the proline-rich N-terminal arm. This last point might help explain the many contradictory data that currently exist concerning the specific intracellular localisation of LITAF-domain-containing proteins in a variety of cell types [9–11, 13, 18].
Consistent with a protein domain that resides in close proximity to membranes, we also show that specific residues within the LITAF domain interact with PE head groups. PE is the second most abundant phospholipid (behind PC) in mammalian cells, accounting for, on average, approximately 25% of cellular phospholipids. PE is found across all intracellular membranes and, as a consequence of its cone-shape, has been proposed to play an essential role in membrane fusion and curvature . By accommodating and interacting with PE head groups, it is tempting to speculate that LITAF domains might therefore play further roles in the regulation of membrane invagination and tubulation, which are essential for the sorting and recycling of membranous cargoes on endosomes .
Our data are consistent with LITAF being a monotopic membrane protein anchored to membranes via an in-plane helical membrane anchor, present within the LITAF domain . However, efforts to analyse the full length LITAF protein, or the predicted hydrophobic helical anchor region in isolation, either by solution NMR or crystallography, were unsuccessful, likely due to the inability of the tested detergent and lipid micelles to mimic the membrane environment found intracellularly. Furthermore, the molecular weight of LITAF remains too small for current cryo-EM techniques . This again highlights the general issue in structural biology that, despite the fact that membrane proteins make up approximately 30% of all eukaryotic cellular proteins  and constitute around 60% of current drug targets , membrane proteins still account for less than 1% of structures in the Protein Data Bank . Moreover, none of these current membrane protein structures contained within the database demonstrate in-plane membrane anchoring domains as we propose for LITAF. In the absence of a predicted helix-loop-helix ‘hairpin’ motif that inserts into the lipid bilayer as postulated for other integral membrane proteins , we can therefore only speculate at present on how precisely the predicted hydrophobic helical anchor associates with membranes. However, our data regarding the loss of membrane association following the substitution of eight hydrophobic amino acids within the predicted helix, in conjunction with our structural data, lead us to speculate that this anchoring-region embeds into the cytosolic-facing monolayer of the membrane bilayer by adopting an amphipathic character [52, 53].
Schwann cells are predicted to be the principal site of pathology in the demyelinating subtypes of CMT. Due to the significant number of genes associated with demyelinating CMT that are predicted or known to encode proteins that function on endocytic pathways , the critical importance of intracellular membrane trafficking in maintaining peripheral nerve myelination by Schwann cells has been brought to the fore. Most mutations in these demyelinating CMT genes associated with membrane trafficking cause disease in an autosomal recessive manner, pointing towards loss of cellular function as the underlying pathogenic mechanism in these instances. In contrast, CMT1C associated with mutations in LITAF is inherited in an autosomal dominant pattern. Previous reports have confirmed that the expression of LITAF is not critical for peripheral nerve myelination in mice , making the ‘dominant-negative’ hypothesis, due to the possible formation of non-functional LITAF hetero-oligomers (containing wild type and mutant species), less likely. On the contrary, a more probable pathogenic mechanism involves a ‘toxic gain-of-function’ resulting from the altered molecular and cellular properties of mutant LITAF when expressed in vivo . Significantly, the LITAF protein is known to be widely expressed across tissues , suggesting that, while compensatory mechanisms must exist in most cell types, the intracellular membrane trafficking pathways in Schwann cells must be distinctly susceptible to the presence of a mutant copy of LITAF.
But how do CMT1C point mutations cause toxicity? Thus far, all proven CMT1C-associated pathogenic mutations in LITAF are found in the C-terminal LITAF domain and previous reports have suggested that these mutations might disrupt the association of LITAF with membranes , although the precise mechanism remained unclear. Here, we show that the CMT1C-associated pathogenic V144M mutation in LITAF leads to chemical shift perturbations across the LITAF domain that sits in close approximation to the surface of membranes, whilst maintaining an overall secondary structure similar to the wild type protein. However, when the V144M construct was incubated with PE head groups, the disease-associated protein became misfolded, leading to precipitation in vitro, in complete contrast to wild type LITAF. Furthermore, cycloheximide chase experiments in vivo suggested that the V144M construct was degraded more rapidly compared to wild type. It is therefore tempting to speculate that all CMT1C mutations might lead to similar structural alterations within the LITAF domain, resulting in an unstable and toxic interaction with lipid head groups and consequent endocytic membrane disruption [10, 56]. This aberrant association of LITAF with membranes might lead to the dysfunctional trafficking of Schwann cell membrane receptors, including growth factor receptors [9, 57, 58] and cell adhesion molecules [6, 59], with consequent disruption of the critical endocytic pathways that maintain the integrity of peripheral nerve myelin. Such mistrafficking would, in turn, lead to instability of the myelin sheath, demyelination, secondary axonal degeneration and, eventually, the motor and sensory deficits that characterise patients affected by a diagnosis of CMT1C.
In summary, we present the first structural protein model of LITAF based on experimental data, and describe the biophysical and biochemical consequences of a CMT1C-associated pathogenic mutation. We anticipate that this work will form a platform for the future study of LITAF, and LITAF-domains in general, and their role in intracellular membrane trafficking. In the absence of any current treatments for CMT, we believe that such investigation will not only lead to the identification of future therapeutic strategies for CMT1C, but will also highlight molecular and cellular dysfunctional pathways common to more than one subtype of this common, currently incurable, neurological disorder.
Reagents and cell culture
Antibodies and reagents used during this study include mouse anti-LITAF (BD Biosciences, #611614, RRID: AB_399056, Lot #29025), rabbit anti-VPS26 (gift from M. N. J. Seaman, Cambridge UK), mouse anti-LAMP1 (DSHB, H4A3, RRID: AB_2296838), mouse anti-EEA1 (BD Biosciences, #610457, RRID: AB_397830, Lot #5023919), rabbit anti-TGN46 (Abcam, ab50595, RRID: AB_2203289, Lot #GR133666-6), mouse anti-HA (16B12, Covance, mms-101R-50), anti-myc (9B11, Cell Signaling, #2276S, RRID: AB_10693333, Lot #19), rabbit anti-Flag (Cell Signaling, #2368, RRID: AB_2217020, Lot #6), rabbit anti-calnexin (Santa Cruz, sc-11397, RRID: AB_2243890, Lot #F1808), rabbit anti-NEDD4 (Abcam, ab14592, RRID: AB_301364), mouse anti-GAPDH (Sigma, G8795, RRID: AB_1078991, Lot #092M4820V), mouse anti-β-lactamase (Abcam, ab12251, RRID: AB_298974), mouse anti-α-tubulin (DSHB, AA4.3, RRID: AB_579793, Lot #8/29/13), rabbit anti-GFP (Thermo Fisher Scientific, A11122, RRID: AB_10073917, Lot #1453341), CF488A Wheat Germ Agglutinin (Biotium, #29022-1, Lot #15 W0225), Alexa Fluor 488- (Thermo Fisher Scientific, A11034, RRID: AB_2576217, Lot #939304), Alexa Fluor 568-conjugated goat anti-rabbit H + L (Thermo Fisher Scientific, A11036, RRID: AB_10563566, Lot # 757102), goat anti-mouse H + L (Thermo Fisher Scientific, A11031, RRID: AB_144696, Lot #1126619), Alexa Fluor 488-conjugated goat anti mouse IgG1 (Thermo Fisher Scientific, A21121, RRID: AB_2535764, Lot #1485220), and Alexa-Fluor 555-conjugated anti-mouse IgG2a (Thermo Fisher Scientific, A21137, RRID: AB_2535776, Lot # 1541489). HRP-conjugated goat anti-rabbit and goat anti-mouse antibodies were used as secondary antibodies for western blotting (Sigma-Aldrich). Proteins separated by SDS-PAGE followed by transfer to nitrocellulose membranes were detected using WesternBright ECL western blotting detection kit (Advansta, CA, USA).
Full length cDNAs for LITAF (IMAGE clone 4000250) and NEDD4 (IMAGE clone 8860509) were obtained from Source Bioscience (Cambridge, UK). LITAF Δ114–139 and LITAF N-helix constructs were generated using synthesized gBlocks (Integrated DNA Technologies, Iowa, USA) and Gibson cloning into the relevant expression vectors (NEB) . Site-directed mutagenesis was performed using a QuikChange II XL kit (Stratagene). All DNA constructs were sequenced and validated by Source Bioscience (Cambridge, UK).
HeLa cells (gift from F. Buss, Cambridge UK) were grown at 37 °C in RPMI (Sigma-Aldrich) containing 10% FCS and 2 mM L-glutamine, 100 U/mL penicillin and 100 μg/mL streptomycin in a 5% CO2 humidified atmosphere. RPE cells (ARPE-19, ATCC) were grown at 37 °C in DMEM:Ham’s F-12 (50:50) (Sigma-Aldrich) containing 10% FCS and 2 mM L-glutamine, 100 U/mL penicillin and 100 μg/mL streptomycin in a 5% CO2 humidified atmosphere. Cells were transfected using polyethylenimine (Polysciences, PA, USA) or Lipofectamine LTX (Thermo Fisher Scientific).
Comparative genomics and phylogenetics
Characterised LITAF domain protein sequences from H. sapiens were used as initial queries to identify putative LITAF domain proteins from the genomes of the organisms listed in (Additional file 1: Figure S1). Where no homologue was identified, secondary searches were carried out using identified sequences from more closely related taxa. Genome searches were carried out using the basic local alignment search tool (BLAST) against publicly available genome databases accessible via NCBI  with exception of those listed below; Plasmodium falciparum (PlasmoDB, ); Toxoplasma gondii (ToxoDB, ); Naegleria gruberi, Guillardia theta, Emiliania huxleyi, Bigelowiella natans, Phaeodactylum tricornutum, Phytophthora sojae, Psuedo nitzschia multiseries, Populus trichocarpa, Selaginella moellendorffii, and Chlamydomonas reinhardtii (Joint Genome Initiative, US Department of Energy ); Bodo saltans (GeneDB ); Leishmania major, Trypanosoma cruzi, and Trypanosoma brucei (TriTrypDB ); and Giardia lamblia (GiardiaDB ). All identified LITAF domain sequences were verified by reciprocal BLAST against the non-redundant protein database (NCBI). For phylogenetic reconstruction, protein sequences were aligned using MAFFT via jalview [68, 69] and edited manually to remove gaps and poorly conserved regions. Phylogenies were reconstructed using Bayesian (MrBayes) and bootstrapped maximum likelihood (RaxML, PhyML) approaches. PhyML was run through the South of France Bioinformatics Platform web server . RaxML and MrBayes were run through the Cyberinfrastructure for Phylogenetic Research (CIPRES) Science Gateway web server . MrBayes version 3.2.2 analyses were run using a mixed model for 1 × 106 generations, with convergence verified by standard deviation of split frequencies < 0.05. All trees before plateau were removed as burn-in. The appropriate evolutionary model was assigned using protest v2.4 . Domain prediction was carried out via the NCBI conserved domain database (CDD), HMMScan (HMMER, Janelia)  and InterProScan .
For general immunofluorescence microscopy, cells were plated on glass coverslips, fixed in 4% paraformaldehyde and permeabilised with 0.1% TritonX-100 in phosphate buffered saline (PBS) at pH 7.4. Bovine serum albumin (BSA; 1%) in PBS was used to block fixed cells before incubation with primary antibodies followed by AlexaFluor-conjugated secondary antibodies. Cover slips were mounted onto glass slides using ProLong Gold Antifade Mountant with DAPI (Thermo Fisher Scientific). Wide-field fluorescent images were obtained using a Zeiss Axioimager Z2 Upright Wide-field microscope (Carl Zeiss), and confocal images were acquired using Zeiss LSM710 and LSM880 confocal microscopes (Carl Zeiss). Imaging data were analysed using Zeiss ZEN software (Carl Zeiss) and Volocity (PerkinElmer).
Membrane fractionation and TX114 extraction
To isolate cellular membranes, HeLa cells were washed with cold 10 mM Tris pH 7.4 before scraping into a small volume of 10 mM Tris pH 7.4, 10 mM NaCl and 1.5 mM MgCl2. The cells were then lysed by gently passing through a 25G needle multiple times before centrifugation at 1330 g for 10 minutes. The resulting supernatant was then sonicated briefly on ice before further centrifugation at 280,000 g for 45 minutes to obtain a membrane fraction (pellet) and cytosolic supernatant. The membranes were resuspended in an equal volume of buffer to the supernatant before proceeding to SDS-PAGE and western blotting. For the LITAF cysteine mutants, the membrane pellets were additionally incubated with 10 mM Tris pH 7.4, 10 mM NaCl, 1.5 mM MgCl2 and 1% TX100, and complete protease inhibitor (Roche) to extract soluble membrane proteins before a second centrifugation at 280,000 g for 45 minutes. The soluble supernatant was transferred to a new tube and the insoluble pellet was resuspended with an equal volume of 10 mM Tris pH 7.4, 10 mM NaCl and 1.5 mM MgCl2 containing protease inhibitors before proceeding to SDS-PAGE and western blotting. Phase partitioning of endogenous LITAF and marker protein from HeLa cell membranes with Triton X114 (Sigma-Aldrich) was performed as previously described , with equal proportions of the detergent and aqueous phases being analysed by western blotting.
Determination of LITAF membrane topology
HeLa cells stably expressing HA-LITAF-myc were generated using the pLXIN retroviral system (Clontech) as previously described , and a single transduced clone selected for further study to ensure consistency across experiments. Cells were seeded onto glass cover slips and left to settle overnight. The following steps until fixation with paraformaldehyde were performed on ice at 4 °C. The cells were first washed twice with ice-cold KHM buffer (20 mM HEPES pH 7.4, 110 mM potassium acetate, 2 mM MgCl2). Digitonin (20 μM; Sigma-Aldrich) was then used to permeabilise the plasma membrane by incubating for 3 minutes on ice before washing three times with ice-cold KHM buffer. Free aldehydes where then quenched with 50 mM NH4Cl in PBS for 5 minutes before twice washing with PBS. Cells were next blocked with 1% BSA in PBS for 30 minutes before incubation with primary antibodies for 1 hour on ice at 4 °C. Following three further washes with PBS, the cells were fixed with 4% paraformaldehyde at room temperature for 15 minutes, washed with PBS and incubated with AlexaFluor-conjugated secondary antibodies before mounting on glass slides for immunofluorescence microscopy. Digitonin (100 μM) was used to permeabilise both endosomal membranes and plasma membranes, additionally exposing epitopes on the luminal side of endocytic vesicles.
For cell-free expression of HA-LITAF-myc, a rabbit reticulocyte lysate cell-free expression system was used in the presence of canine microsomal membranes (Promega). The signal peptide-containing protein, β-lactamase, served as a control for membrane insertion and translocation into the lumen of microsomes, and hence protection from added protease digestion. The expressed proteins were incubated with proteinase K (NEB) for 1 hour at 37 °C before separation by SDS-PAGE followed by western blotting.
Bacterial protein expression and purification
cDNAs corresponding to wild type LITAF, Δ114–139 and N-helix were cloned into the pOPINS vector before expression of the resulting His6-SUMO-tagged protein constructs in C43(DE3) BL21 E. coli. Bacterial lysates were prepared by cell disruption of the bacterial pellet in lysis buffer (50 mM HEPES pH 7.4, 300 mM NaCl, 3 mM 2-mercaptoethanol, 200 μM AEBSF) followed by centrifugation of insoluble material at 100,000 g for 30 minutes. His6-SUMO-LITAF constructs were purified from lysates using Ni-NTA resin (Qiagen) and the eluting imidazole removed using Centripure P25 Zetadex gel filtration columns (Generon) and the protein buffer-exchanged into 20 mM HEPES pH 7.4, 100 mM NaCl, 0.5 mM TCEP. The purified protein was then incubated with ULP1 SUMO protease for 1 hour at room temperature and the His6-SUMO tag removed by a second incubation with Ni2+-NTA. The final purification step consisted of size exclusion chromatography using a Superdex 200 10/300 gel filtration column (GE Healthcare Life Sciences). Protein samples were concentrated using Vivaspin centrifugal concentrators (Sartorius, UK) and any insoluble particles removed using Proteus clarification mini spin columns (Generon). For NMR analysis, proteins were expressed in C43(DE3) BL21 E. coli grown in minimal media containing ammonium-15N chloride (Sigma-Aldrich) and/or 13C-D-Glucose (Cambridge Isotopes) as previously described . For wild type LITAF constructs (containing the predicted hydrophobic helical membrane anchor), detergents were required at each step to maintain solubility. For the experiments described in this study, 1% DDM (w/v) or 1% FC12 were used for cell lysis and 0.04% DDM (w/v) or 0.125% FC12 added to each subsequent buffer when dealing with the full length wild type LITAF protein. For GST-tagged proteins, cDNA encoding residues 1–77 of human LITAF and residues 596–944 of human NEDD4 were cloned into pGEX-4 T1 expression vector (GE Healthcare Life Sciences) for expression in C43(DE3) BL21 E. coli. Purification was performed using glutathione sepharose 4B according to the manufacturer’s instructions (GE Healthcare Life Sciences). For NMR analysis, purified GST-NEDD4 596–944 was buffer-exchanged into 20 mM HEPES pH 7.4, 100 mM NaCl and 0.5 mM TCEP by using a Superdex 75 10/300 gel filtration column (GE Healthcare Life Sciences).
GST pull-down assays
Cell lysates were prepared by scraping RPE cells into a small volume of lysis buffer (20 mM HEPES pH 7.4; 150 mM NaCl; 1 mM EDTA; 0.5% Triton X-100 (v/v); complete protease inhibitor (Roche)) and transferring into a 1.5 mL microcentrifuge tube. The lysate was then sheared by gently passing the solution back and forth through a 21-G needle followed by a 25-G needle, before centrifugation at 20,000 g for 30 min at 4 °C. The resulting supernatant was first precleared by gentle agitation for an hour at 4 °C with glutathione sepharose 4B beads (GE Healthcare Life Sciences) before the addition of GST-LITAF 1–77 constructs at a concentration of 0.2 mg/mL. Following a further incubation for 2 hours at 4 °C with gentle agitation, glutathione sepharose 4B beads were added and incubated for 1 hour more with gentle agitation at 4 °C. The beads were washed twice with lysis buffer and once with PBS before elution of the bound proteins by heating in SDS sample buffer (6 M Urea, 1% SDS (w/v), 1 M 2-mercaptoethanol and 150 mM Tris (pH 6.7)). Eluted proteins were separated by SDS-PAGE and transferred to nitrocellulose membranes for western blotting.
RPE cells stably expressing wild type HA-LITAF or HA-LITAF V144M were seeded before incubation with 300 μg/mL of cycloheximide (Sigma). Cells were lysed in RIPA buffer at 0, 1, 2, 4, 8 and 24 hours after incubation with cycloheximide before brief sonication and centrifugation at 20,000 g for 30 minutes at 4 °C. The supernatants were transferred to new tubes and the protein concentration determined using Precision Red reagent (Cytoskeleton, Inc.). Equal amounts of protein were separated by SDS-PAGE before analysis by western blotting.
The unambiguous identity of the metals in the recombinantly expressed LITAF proteins and their stoichiometry was determined using microPIXE (microbeam Proton Induced X-ray Emission), an established technique for such analyses . The recombinant proteins were prepared as described above before buffer exchange into 10 mM Tris pH 7.4, 200 mM NaBr and 0.5 mM TCEP using a Superdex 75 GL size exclusion chromatography column (GE Healthcare Life Sciences). DDM (0.04%) was also used to maintain the solubility of wild type LITAF constructs for the purpose of these experiments. Resulting protein concentrations, estimated by measuring the absorbance at 280 nm, were 1.7 mg/mL, 4.7 mg/mL and 1.7 mg/mL for wild type LITAF, LITAF Δ114–139 and LITAF N-helix, respectively. This pre-measurement buffer exchange was also necessary to remove any traces of sulfur and chlorine. For micro-PIXE measurements of proteins, sulfur acts as an internal standard, and because of the proximity of the X-ray emission energies of sulfur and chlorine to one another, strong chloride fluorescence can affect the accuracy with which the sulfur peak can be quantified . The measurements were carried out at the Ion Beam Centre, University of Surrey, UK . A 2.5-MeV proton beam of 3 μm in diameter was used to induce characteristic X-ray emission from dried liquid protein droplets (volume per droplet = 0.1 μL) under vacuum. The X-rays were detected in a solid state lithium drifted silicon detector with high energy resolution. By scanning the proton beam in x and y over the droplet, spatial maps were obtained of all elements heavier than magnesium present in the sample. Quantitative information was obtained by collecting 3- or 4-point spectra from each droplet. These spectra were analyzed with GUPIX  within DAN32  to extract the relative amount of each element of interest in the sample.
NMR experiments and analysis
15N- and 13C-labelled proteins were prepared in 20 mM HEPES pH 7.4, 100 mM NaCl, 0.5 mM Tris(2-carboxyethyl)phosphine hydrochloride (TCEP) at typical concentrations of between 60 and 250 μM. All experiments were collected at 298 K on Bruker Avance III 600 MHz and Avance 2+ 700 MHz spectrometers equipped with cryogenic triple resonance TCI probes. Data were processed using the software packages Topspin 3.2 (Bruker) and both Multi-Dimensional Decomposition  and Compressed Sensing  for data collected by Non Uniform Sampling. All NMR data were analyzed using SPARKY 3 (T. D. Goddard and D. G. Kneller, University of California, San Francisco).
1D proton transverse relaxation (T2) experiments were used to estimate the molecular weight of the LITAF domain. For a globular protein, its transverse relaxation is governed by its molecular weight by the following relationship: MW × 0.5 ≈ 1/5 × T2, where T2 is in seconds and MW is in kDa. Standard 1D proton transverse relaxation experiments were collected with different delays (1 and 2), with observed peak intensities reduced after an increased delay. T2 can therefore be estimated using the following equation: T2 = 2(Δ1–Δ2)/ln(I1/I2), where I is the peak intensity and Δ is the relaxation delay.
Backbone chemical shift assignments were completed using HNCO, HNCACO, HNCA, HNCACB and CBCACONH. The assignment of proline residues was aided by 13C observed CON experiments complimented with HCBCANCO, HCBCACON, HNCANNH and HNCOCANNH experiments. Side chain HA and HB were assigned using an HBHACONH experiment. Through space contacts were established by 15N and 13C HSQC NOESY experiments.
Secondary structure predictions were made using N, HN, CA, CB and HA chemical shifts and TALOS+ .
Assigned BEST-TROSY (band selective excitation short transients transverse relaxation optimized spectroscopy) were used to make weighted chemical shift maps on mutation and head group or NEDD4 WW domain interactions using the equation ((Δ1H)2 + (Δ15N/5)2)0.5, where the Δ denotes the difference in parts per million of the chemical shift between the assigned apo peak and its nearest neighbour in the mixed sample spectra.
In the head group experiments, PG, PC, PS, PE and IP6 head groups were sourced from Sigma-Aldrich and incubated with LITAF (60 μM concentration) at molar ratios of 1:1000 (PG, PC, PE) and 1:500 (PS, IP6). The pH of each LITAF/head group mixture was checked and found to be the same as the sample used to generate the control spectra.
LITAF structure modelling
NMR assignment information was submitted to the CS Rosetta webserver for de novo structure calculations [31, 84], which did not return a convergence of structures, suggesting a low confidence in the prediction. Given the established metal coordination of the LITAF constructs and NMR restraints, a zinc-bound structure model methodology was then followed using CS-Rosetta fragment files with the abinitio.metalrelax protocol of the Rosetta package . The jump restraint file was edited so that the zinc atom coordinated cysteine residues C96, C99, C148 and C151 (full length residue numbers). Running the scripts for both the LITAF Δ114–139 and wild type LITAF domains returned 5000 structural models for each construct (also referred to as decoys) of improved confidence. These decoys were clustered with a 5 Å cutoff using Calibur  and scored for similarity to the centre of each cluster using the score_jd2 script that is part of the Rosetta package. The structures shown in Figs. 6 and 8 were representative of the lowest energy structures of the largest cluster in each calculation round.
We thank Professor J Paul Luzio for support and advice. We acknowledge Matthew Gratian and Mark Bowen (Cambridge Institute for Medical Research) for technical assistance with microscopy and image analysis, and thank Dr Geoff Grime at the Surrey Ion Beam Centre for assisting with the microPIXE experiments.
This work was supported by a Wellcome-Beit Prize and Intermediate Clinical Fellowship to RCR (093809/Z/10/Z). The Cambridge Institute for Medical Research (CIMR) is supported by Wellcome Trust Strategic Award 100140, and the Zeiss LSM710 confocal microscope was purchased with Wellcome Trust grant 093026.
Availability of data and materials
All NMR data have been deposited in the Biological Magnetic Resonance Bank (BMRB) with accession number 26940 (http://www.bmrb.wisc.edu/). The NMR-based structural models are available from the corresponding author by request.
AKH, JLW, LW, SQ, SMVF and RCR performed the experiments. PTM performed the evolutionary sequence analysis. EFG performed the microPIXE experiments. RCR wrote the manuscript, which was edited by all the co-authors. All authors read and approved the final manuscript.
The authors declare that they have no competing interests.
Consent for publication
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