A functional chaperone condensate in the endoplasmic reticulum

doi:10.21203/rs.3.rs-4796355/v1

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A functional chaperone condensate in the endoplasmic reticulum

https://doi.org/10.21203/rs.3.rs-4796355/v1

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One third of eukaryotic proteins are processed within the endoplasmic reticulum (ER) to obtain their correct structure^1-3. This function is ensured by a network of molecular chaperones that recognizes client proteins and assists their folding^4,5. How the ER chaperones organize in a supramolecular manner to exert their cooperativity has, however, remained unclear. Here, we report the discovery of a multi-chaperone condensate in the ER lumen, which is formed around the chaperone PDIA6 during protein folding homeostasis. We resolve the structure of PDIA6 and the mechanism underlying its condensate formation at the atomic and cellular level. We find that the multivalency required for PDIA6 phase separation is created by two specific interfaces. One of these interfaces is regulated by a Ca²⁺-dependent molecular switch, allowing dynamic adaptations to ER stress. The PDIA6 condensates recruit further chaperones, in particular Hsp70 BiP, J-domain protein ERdj3, disulfide isomerase PDIA1 and Hsp90 Grp94, which are the essential components of the early folding machinery. The chaperone condensates enhance folding of client proteins, which we show exemplarily on proinsulin, and prevent protein misfolding in the ER lumen. The PDIA6-scaffolded chaperone condensates hence provide the functional basis for spatial and temporal coordination of the dynamic ER chaperone network.

Biological sciences/Cell biology/Protein folding/Endoplasmic reticulum

Biological sciences/Structural biology/NMR spectroscopy/Solution-state NMR

Biological sciences/Molecular biology/Protein folding/Chaperones

Molecular chaperones

protein folding

endoplasmic reticulum

liquid-liquid phase separation

condensate

ER-stress

protein disulfide isomerase

PDIA6

Hsp70 BiP

insulin

The protein disulfide isomerase (PDI) chaperone family is indispensable for the maintenance of ER protein folding homeostasis^6,7. While all PDIs share common structural features, they differ in terms of client protein specificity and affinity, chaperone activity and redox properties, resulting in specialized roles for individual family members⁸. One particular family member, PDIA6, caught our attention due to its direct connection to several human diseases including cancer and diabetes, yet its exact functioning remained elusive^9-12.

PDIA6 is organized in phase-separated condensates

We observed that endogenous PDIA6 localized in the ER in human HeLa cells, as expected (Fig. 1a). Remarkably, however, the endogenous protein was not homogeneously dispersed, but concentrated locally in well-defined clusters. Overexpressed PDIA6 formed clusters of similar shape and localization in living cells, albeit with substantially increased size (Fig. 1a and Extended Data Fig. 1a-c). PDIA6 formed similar clusters also in HEK and U2OS cells (Extended Data Fig. 1 d,e). The PDIA6 clusters were mobile within the ER and underwent fission and fusion events (Fig. 1b and Supplementary Videos 1,2). Their fluorescence recovery after photobleaching (FRAP) occurred on the timescale of minutes, which is common for biological phase separated condensates^13,14 (Fig. 1c and Supplementary Video 3). The PDIA6 condensates thus fulfill all hallmarks of biological phase separation¹⁵.

PDIA6 condensates disperse in response to ER stress

Malfunction of PDIA6 has been reported to lead to an imbalance in ER folding homeostasis. We thus wondered whether a functional coupling exists between ER stress and condensate formation. To create a physiological ER stress situation, we overexpressed Akita proinsulin, a misfolded mutant of proinsulin that accumulates in the ER and thereby causes ER stress¹⁶. This led to homogenous dispersion of PDIA6 in the entire ER lumen (Extended Data Fig. 2a). Similarly, upon treatment of the cells with the pharmacological ER stress inducer tunicamycin (Tm) the PDIA6 condensates dissolved and PDIA6 dispersed, independently of its expression levels (Extended Data Fig. 2b-d). The stress-dependent condensate dissolution was reversible, as PDIA6 condensates reorganized after washout of the stressor. Two other stress inducers, thapsigargin (Tg) and cyclopiazonic acid (CPA), induced the same effect (Extended Data Fig. 2e,f). Remarkably, however, the dissolution kinetics substantially differed between the compounds. PDIA6 condensates dissolved completely within 1 h upon Tg treatment, while dissolution was observed only after 6 h of Tm treatment (Fig. 1d,e and Extended Data Fig. 2g). This kinetic difference correlates with the different modes of action of the two drugs. Tg blocks the SERCA Ca²⁺ pumps¹⁷, leading to an almost immediate Ca²⁺ depletion in the ER lumen, whereas Tm inhibits N-glycosylation of newly synthesized proteins¹⁸ and causes a delayed drop in Ca²⁺ levels as a secondary effect of the accumulation of unfolded proteins¹⁹. Overall, formation of PDIA6 condensates is thus coupled to protein homeostatic conditions and their dissolution in response to ER stress regulated directly by luminal Ca²⁺ levels.

To test this mechanistic hypothesis, we reconstituted droplets of purified recombinant PDIA6 protein in vitro. Homeostatic ER was mimicked by high Ca²⁺ concentration, reducing conditions, physiological pH and a crowding agent. Under these conditions, recombinant PDIA6 reproducibly formed droplets that underwent fusion events and time-dependent growth on the timescale of seconds. The protein dynamically exchanged between the condensed and dispersed phase with similar kinetics as in vivo, as determined by FRAP experiments (Fig. 1f,g and Extended Data Fig. 2h-k). Droplet formation was not induced by another divalent ion, Mg²⁺, i.e. it was Ca²⁺-specific. Importantly, the droplets formed only at Ca²⁺ concentrations above ~500 μM (Fig. 1h), which corresponds to the physiological transition between protein folding homeostasis ([Ca²⁺]_homeo ≈ 800 μM) and ER stress ([Ca²⁺]_stress ≈ 100 μM)²⁰, and therefore correlates very well with our observations in living cells. PDIA6 condensates are thus regulated by the Ca²⁺ concentration.

Structural characterization and domain organization of PDIA6

We then employed an integrated structural biology approach to resolve the structure and mechanism of PDIA6 condensate formation. PDIA6 consists of two catalytically active thioredoxin-like domains, domain a⁰ and a, which are followed by the inactive domain b of unknown function (Fig. 2a)^21-22. In order to monitor conformational changes and interactions by NMR spectroscopy, we assigned 94% of the Met^e, Leu^δ2 and Val^γ2 methyl groups and established near-complete amide backbone assignments (Extended Data Fig. 3). Using SEC-MALS and NOESY experiments, we found that PDIA6 dimerizes via helix α4 in domain a⁰ (Fig. 2b,c and Extended Data Fig. 4a), in agreement with previous findings²³. The dimerization interface was resolved by a methyl NOE network between residues L72 and L134, as well as between V75 and L131, L134, L135, which corresponds to the arrangement observed in crystals of domain a⁰ (PDB: 4ef0). The dimerization thus resulted from a C-terminal extension of helix α4 in domain a⁰ by seven residues compared to the other characterized monomeric PDI family members PDIA1-5²⁴ (Extended Data Fig. 4b,c). Dimerization occurred with a dissociation constant in the low nanomolar range (Extended Data Fig. 4d), and the dimer is therefore the dominant species at the physiological concentration of around 100 μM^25,26. We also determined the crystal structure of domainb (residues 274–427), which adopts a thioredoxin-like fold, to a resolution of 1.8 Å (Extended Data Fig. 4e,f and Supplementary Table 1).

The dimeric domain a⁰is connected to domain a by a long, positively charged linker, and domains a and b are connected by a short 5-residue linker that restricts the relative domain motions by three proline residues (Figure 2d). Domains a and b thus interact strongly, as confirmed by large chemical shift perturbations (CSP) upon separation and an interdomain NOE cross peak (Extended Data Fig. 4g-i). The C-terminal tail of PDIA6 (residues 420–440) is highly flexible and disordered, as evidenced by a narrow chemical shift dispersion and high NMR signal intensities (Fig. 2e). Combining our experimental data, we established a full atomic model of PDIA6 (Fig. 2f).

PDIA6 has three distinct Ca²⁺ binding sites

As PDIA6 condensate formation is Ca²⁺-dependent, we set out to determine PDIA6 Ca²⁺ binding sites. Titration of Ca²⁺ itself did not lead to significant changes in the methyl NMR spectra, demonstrating the absence of large structural rearrangements upon Ca²⁺ binding (Extended Data Fig. 5a). Titration of the paramagnetic analog Gd³⁺, however, quenched NMR signals specifically in three structural regions I, II and III, revealing three cation binding sites (Fig. 2g,h and Extended Data Fig. 5b,c). Bound Gd³⁺ was outcompeted by an excess of Ca²⁺, proving Ca²⁺ binding specificity. Binding sites I and II are located at the domain a/b interface and in domain b, respectively. Binding site III is located within the disordered C-terminal tail, as previously proposed²³. We pinpointed the exact locations of binding sites I and II in the center of the quenched regions by structural alignment with the Ca²⁺binding protein calsequestrin (Fig. 2i and Extended Data Fig. 5d,e)²⁷. Based on the competition titrations, we estimated the dissociation constant of Ca²⁺ to be in the range of 100–1000 μM, which is consistent with the Ca²⁺-dependent condensate formation switch that we observed in vitro and in vivo. Ca²⁺ binding thus activates a molecular switch that controls PDIA6 phase separation.

A molecular switch in domain b regulates PDIA6 condensate formation

Next, we set out to determine the molecular mechanism underlying condensate formation. Biological phase separation relies on multivalent interactions¹⁵, and therefore, we expected at least two intermolecular interactions to be involved. We suspected that dimerization of domain a⁰ is one of these interactions and indeed, a monomeric construct of PDIA6²³ did not form condensates (Extended Data Fig. 6a). In search for the second interaction, we first tested droplet formation of the individual domain subconstructs, but none of these formed droplets in solution. We also probed a potential role of the disordered C-terminal tail, but neither neutralization of its positive charges (EQDN), nor its entire truncation (ΔC) abrogated the formation of condensates (Extended Data Fig. 6b,c).

We hypothesized that the highly flexible and charged linker connecting domains a⁰ and a (Fig. 3a) might drive condensate formation by interacting with another PDIA6 dimer. To resolve this interaction, we employed NMR spectroscopy on the disperse protein, i.e. in absence of crowding agent. We suspected this interaction to be rather weak and therefore added a 21-residue peptide corresponding to the a⁰–a linker (GGRSGGYSSGKQGRSDSSSKK) to methyl-labeled full length PDIA6 in order to saturate any potential binding sites. Indeed, the addition of the peptide led to drastic spectral changes in domain b. In absence of the peptide, the domain equally populated two conformations in slow exchange (2 peaks/residue for all residues in domain b) (57:43 ± 8 %). Upon titration of the linker-peptide this equilibrium strongly shifted towards one conformation (93:7 ± 3%), evidencing a transient interaction between the linker and domain b (Fig. 3b,c and Extended Data Fig. 6d-g). High salt concentration as well as charge-neutralizing mutations in the linker-peptide impaired the interaction, validating its electrostatic character (Fig. 3c,d and Extended Data Fig. 6h-j). Strikingly, the interaction was Ca²⁺-dependent, in full agreement with the physiological observation of phase separation.

The data thus suggest a functional model for PDIA6 condensate formation, where the positively charged a⁰–a linker dynamically interacts with the negatively charged surface in domain b at homeostatic Ca²⁺ concentrations. We tested this functional model on the in vitro reconstituted droplets of full-length PDIA6. In full agreement with the model, either increased salt concentration or neutralization of the positively charged linker residues within the protein by mutagenesis prevented droplet formation (Fig. 3e and Extended Data Fig. 6k,l). Encouraged by these findings, we then also probed the model in cells. Neutralization of the positively charges in the linker should abolish PDIA6 condensate formation. Indeed, cells transfected with PDIA6 linker mutants no longer formed the large condensates observed for PDIA6 WT. Removal of two charges from the linker led to a reduction in condensate size and complete neutralization of all five charged residues (5x mutant) fully impeded condensate formation, acting as dominant negative (Fig. 3f). The expression levels of endogenous PDIA6 and transfected 5x mutant were approximately equal in our in vivo experiment, and at the same stoichiometry PDIA6 droplet formation was equally abrogated in vitro (Extended Data Fig. 6m,n). The multivalency required for PDIA6 condensate formation thus results from (i) the high-affinity dimerization of domain a⁰ and (ii) the transient, Ca²⁺-dependent electrostatic interaction between the a⁰–a linker and domain b (Fig. 3g-i).

PDIA6 recruits other chaperones into the condensates

To explore the biological role of the PDIA6 condensates, in particular their functional role in the ER chaperone network, we probed the localization of the central chaperone Hsp70 BiP²⁸ relative to the condensates. BiP is one of the most abundant ER chaperones involved in protein folding and disaggregation and has been reported to interact with PDIA6^29-32. Immunostaining of HeLa cells showed that endogenous BiP colocalized in the PDIA6 condensates, both at endogenous and overexpressed PDIA6 levels (Fig. 4a,b and Table S2). BiP overexpression, in turn, did not affect the PDIA6 condensates (Extended Data Fig. 7a). We also tested whether BiP would form condensates on its own by staining endogenous BiP in cells transfected with PDIA6 5x linker mutant. As shown above, in these cells PDIA6 does not form condensates but is homogenously dispersed throughout the entire ER. Under this condition, BiP did not form condensates, thus establishing PDIA6 as the scaffold of the condensates to which it recruits BiP (Extended Data Fig. 7b).

Next, we employed methyl and amide NMR experiments to determine the interaction sites between PDIA6 and BiP by chemical shift mapping. PDIA6 interacts via its domain b with the nucleotide binding domain (NBD) of BiP (Fig. 4c and Extended Data Fig. 7c-j). The affinity of K_D ≈ 50 μM (Extended Data Fig. 7k) is in the same range as known BiP-co-chaperone interactions^33,34, and fully in line with a dynamic chaperone network. Importantly, the BiP binding site on domain b does not overlap with the interface required for condensate formation, such that condensate formation and BiP recruitment by PDIA6 is simultaneously possible, as observed in cells. In full accordance with this model, the isolated BiP NBD colocalized in PDIA6 droplets in vitro, while it did not undergo phase separation by itself (Fig. 4d). Taken together, these experiments clearly establish that PDIA6 forms the scaffold of the condensates into which it recruits BiP as a secondary protein.

Next, we investigated whether further ER chaperones were also concentrated into PDIA6 condensates. We chose to examine the BiP co-chaperone ERdj3, an ER-resident member of the J-domain protein (JDP) family, because it plays a central role in mediating client protein transfer to BiP³⁵. Strikingly, ERdj3 colocalized with the PDIA6-BiP condensates, both at endogenous protein levels and in PDIA6-transfected cells (Fig. 4e, Extended Data Fig. 8a and Table S2). Next, we probed the localizations of PDIA1, the major protein disulfide isomerase for numerous ER client proteins, and Grp94, the ER-localized Hsp90, which cooperate with BiP in client protein processing^32,36. Strikingly, both chaperones enriched in the PDIA6 condensates at endogenous protein levels and in PDIA6-transfected cells (Fig. 4e, Extended Data Fig. 8a and Table S2). In contrast to this, the lectin chaperones calreticulin and calnexin, responsible for the glycosylation of client proteins prior to their export from the ER³⁷, did not enrich in the PDIA6 condensates (Fig. 4f, Extended Data Fig. 8b,c and Table S2). From these observations, it emerges that the essential components of the early folding machinery – Hsp70, Hsp90, JDPs and PDIs – specifically concentrate in the condensates, whereas the lectin chaperones involved in later stage client maturation – calnexin and calreticulin – are not enriched (Figure 4g).

The chaperone condensates function as folding factories

The colocalization of multiple key chaperones in the condensates strongly suggests a fundamental biological function in client protein folding. Proinsulin, the precursor of insulin, is folded in the ER by the assistance of various chaperones including BiP, Erdj3, PDIA6 and PDIA1^38,39 prior to being exported to the Golgi for further processing and subsequent secretion in form of insulin⁴⁰. This makes it an ideal model client to study the potential role of the condensates as folding compartments.

To assess the spatial localization of proinsulin within the ER, cells were transiently transfected with a proinsulin construct including a C-terminal myc-tag. This allowed to visualize the ER-localized proinsulin by myc-antibody staining and revealed that proinsulin was highly enriched in endogenous PDIA6 condensates (PCC=0.93±0.1) (Fig. 4g,h and Table S2), which strongly points towards a role of the condensates in client protein folding. To evaluate whether proinsulin is indeed actively processed within the PDIA6 condensates, we assessed the effect of the chaperone condensates on insulin secretion. If the PDIA6 condensates served as functional ER folding compartments, their absence should lead to a decrease of insulin secretion. We co-transfected cells with proinsulin and either PDIA6 WT or the PDIA6 5x linker mutant and determined the amount of insulin secretion. Strikingly, the cells transfected with PDIA6 5x linker mutant, which impairs condensate formation, secreted less insulin than PDIA6 WT transfected cells, supporting our hypothesis of a functional role of the PDIA6 condensates in client maturation (Fig. 4i). We would like to point out, that only a fraction of cells was actually co-transfected with PDIA6 and proinsulin, and therefore, the effective difference in insulin secretion is even larger in individual cells. The experiments provide strong evidence that PDIA6 condensates promote folding of insulin.

Finally, we wanted to study the effect of the PDIA6-chaperone condensates on overall ER client protein turnover and proteostasis. To this end, we assessed the effect of chaperone condensates on XBP1 splicing, a well-established readout of the ER folding capacity which directly reports on the accumulation of misfolded proteins in the ER lumen^41,42. We transfected cells with PDIA6 WT or PDIA6 5x linker mutant and quantified sXBP1 levels. Strikingly, the absence of PDIA6 condensates (5x linker mutant) caused a significant increase in XBP1 splicing, directly confirming a functional role of the condensates in overall protein folding (Fig. 4j). It hence seems that the spatial arrangement of PDIA6 in condensates results in more efficient and robust client folding. To test this hypothesis, we subjected mock-transfected, PDIA6 WT or PDIA6 5x linker mutant transfected cells to repeated stresses, with recovery times in between and tested for cell viability. The cell viability of cells expressing the PDIA6 5x linker mutant drastically decreased after repeated stress treatment, compared to PDIA6 WT expressing cells (Extended Data Fig. 8d). The condensates seem to avert cell death in recurring stress situations by preventing the accumulation of misfolded proteins during homeostatic conditions. PDIA6 condensates thus sub-compartmentalize the ER in functional “folding factories”.

In summary, our results reveal that the crucial ER chaperones do not only cooperate in a functional network but form a self-organized supramolecular structure within the ER that guides client folding (Figure 4k). The PDIA6-mediated chaperone condensates appear to act in the early stages of client protein folding and it is thus imaginable that client proteins are imported directly into the condensates upon their translocation into the ER. The physiological benefits of sub-compartmentalizing the ER by arranging chaperones into folding condensates indeed seem to be numerous: The elevated local concentrations of chaperones and clients increases both specificity and folding efficiency, thus reducing protein misfolding and aggregation. The simultaneous decrease in chaperone concentration outside the condensates may prevent non-specific interactions with folded proteins in the ER lumen, thus enabling a multi-step processing pipeline for clients. Furthermore, the dynamic regulation of the chaperone condensates via the Ca²⁺ concentration might help to overcome an increasing load of misfolded proteins in the ER lumen during acute ER stress situations by accelerating protein turnover at the cost of specificity. Condensate properties have recently been reported for the cytosolic chaperones Hsp70 and J-domain proteins, to specifically prevent aggregation of amyloid-forming proteins^13,43. From these and our observations the pattern seems to emerge that sub-compartmentalization by chaperone condensates is a general theme to structure cellular compartments. The chaperone superstructures in the ER not only provide a fascinating layer of functionality to enable protein biogenesis but might also result in novel therapeutical approaches to target protein misfolding diseases.

Protein expression and purification

Cloning, expression and purification of human PDIA6

Human PDIA6 lacking its ER signal sequence (residues 19-440), domain ab (residues 161-440) and domain b (residues 276-440), with an additional N-terminal His₆-Tag including a TEV cleavage sequence and domain a (residues 161-272) with an additional N-terminal His₆-Tag followed by a SUMO tag were synthesis by GenScript. The genes were inserted using NcoI and XhoI restriction enzymes into a pET28a expression vector (Novagen). BL21-(DE3)-Lemo cells (New England Biolabs) were transformed and grown at 37°C in LB medium containing kanamycin (30 mg/mL). Expression was induced at an optical density (OD₆₀₀) of 0.6 by adding 0.5 mM isopropyl-β-d-thiogalactopyranoside (IPTG) at 25 °C for 12 hours. Cells were harvested by centrifugation at 4000 g for 20 min. The pellet was resuspended in 35 mL of lysis buffer per liter of culture (25 mM Hepes (pH 7.5), 300 mM NaCl, 10 mM MgCl₂, 5 mM ATP (Sigma-Aldrich), 0.02 mg/ml ribonuclease, 0.01 mg/mL deoxyribonuclease and 0.2 mg/mL phenylmethylsulfonyl fluoride (PMSF). Cell lysis was performed using a microfluidizer (Microfluidics) for three cycles at 4 °C. The soluble bacterial lysate was separated from cell debris and other components by centrifugation at 14000 g for 60 min and loaded onto a Ni-NTA column (Cytiva) equilibrated in buffer A (25 mM Hepes (pH 7.5), 300 mM NaCl, 10 mM imidazole). PDIA6 eluted at 300 mM imidazole concentration and was dialyzed overnight against buffer (25 mM Hepes (pH 7.5), 300 mM NaCl) to remove the imidazole. PDIA6 was denatured with 6 M urea and loaded onto a Ni-NTA column (Cytiva) equilibrated in buffer A + 6 M urea. PDIA6 eluted at 300 mM imidazole concentration in buffer containing 6 M urea. PDIA6 refolding was achieved by dialysis overnight against buffer (25 mM Hepes (pH 7.5), 300 mM NaCl). After refolding, the His-tag was cleaved by incubation with 1 mg of TEV protease or Ulp1 SUMO protease per 50 mg of PDIA6 in cleavage buffer (25 mM Hepes (pH 7.5), 300 mM NaCl and 1 mM DTT , 0.5 mM EDTA) overnight at 4°C. His-TEV-domain b and his-TEV-domain ab constructs were not cleaved. His-TEV-domain a⁰ and his-TEV domain a used for amide backbone assignment were not cleaved. Cleaved PDIA6 constructs were separated from the TEV or Ulp1 and uncleaved PDIA6 by a reverse Ni-NTA column equilibrated in buffer A. Finally, PDIA6 was concentrated by ultrafiltration and injected to size exclusion chromatography (Superdex-200 16/600 PG, Cytiva or Superdex-75 10/300 PG, Cytiva) to further purify the proteins and adjust the protein to its final buffer (25 mM Hepes (pH 7.5), 150 mM KCl, 10 mM MgCl₂). PDIA6 was concentrated by ultrafiltration and stored at −20 °C until use. Final yield of purified protein was 25 mg per liter of LB medium. To obtain fully reduced PDIA6, the protein was incubated for 1 h at RT with 10 mM TCEP, buffer exchanged (3x) to SEC buffer and immediately frozen. Prior to calcium-binding experiments, PDIA6 was incubated with 20 mM EDTA for 1 h at RT and dialysed against SEC buffer overnight at 4 °C.

Cloning, expression and purification of human BiP

Human BiP lacking its ER signal sequence (residues 19-654) with an N-terminal His₆-Tag including a TEV cleavage sequence was synthesized by GenScript. The gene was inserted through NcoI and XhoI into a pET28a expression vector (Novagen). BL21-(DE3)-Lemo cells (New England Biolabs) were transformed and grown at 37 °C in LB medium. BiP expression was induced at an OD₆₀₀ of 0.6 by adding 1 mM IPTG and expression was continued at 25 °C for 12 hours. Cells were harvested by centrifugation at 5000 g for 20 min. The pellet was resuspended in 20 mL of lysis buffer per liter of culture (25 mM Hepes (pH 7.5), 150 mM NaCl, 10 mM MgCl₂, 0.02 mg/mL ribonuclease, 0.01 mg/mL deoxyribonuclease and 0.2 mg/mL PMSF. Cell lysis was performed using a microfluidizer (Microfluidics) for three cycles at 4 °C. The soluble bacterial lysate was separated from cell debris and other components by centrifugation at 14000 g for 60 min and loaded onto a Ni-NTA column (Cytiva) equilibrated in buffer A (25 mM Hepes (pH 7.5), 150 mM NaCl, 10 mM MgCl₂). BiP eluted at 500 mM imidazole concentration and was dialyzed overnight against buffer (25 mM Hepes (pH 7.5), 300 mM NaCl, 10 mM MgCl₂). BiP was denatured with 6 M urea and loaded onto a Ni-NTA column equilibrated in buffer A + 8 M urea. BiP eluted at 500 mM imidazole concentration in buffer containing 8 M urea. BiP refolding was achieved by dialysis overnight against buffer (25 mM Hepes (pH 7.5), 300 mM NaCl, 10 mM MgCl₂). After refolding, the His-tag was cleaved by incubation with 1 mg of TEV per 50 mg of BiP in cleavage buffer (25 mM Hepes (pH 7.5), 300 mM NaCl, 10 mM MgCl₂ and 1 mM DTT 1 mM, 0.5 mM EDTA) overnight at 4°C. BiP was separated from TEV and uncleaved BiP via a reverse Ni-NTA column (Cytiva) equilibrated in buffer A. BiP was then applied to an anion-exchange column (Cytiva) equilibrated in the buffer QA (25 mM Tris (pH 8.5)) and eluted with 250 mM of KCl. Finally, BiP was concentrated by ultrafiltration and subjected to size exclusion chromatography (Superdex-200 16/600 PG, Cytiva) to further purify the proteins and adjust to its SEC buffer (25 mM Hepes (pH 7.5), 150 mM KCl, 10 mM MgCl₂). Afterward, BiP was concentrated by ultrafiltration and stored at −20°C until use. Final yield of purified protein was 15-20 mg for wild-type BiP per liter of LB medium.

Methyl labeling of human PDIA6 and BiP

Methyl-labeled PDIA6 proteins were obtained by growing the expression cells in M9 minimal media prepared with 99.85% D2O (Sigma-Aldrich) containing ¹⁵NH₄Cl (1 g/L; Sigma-Aldrich) and D-glucose-d₇ (2 g/L; Sigma-Aldrich). At an OD₆₀₀ of 0.8, a solution containing the labeled precursors was added. For [U-²H, ¹⁵N, ¹²C], Met-[¹³CH₃]^ε, Val-[¹³CH₃]^γ2, Leu-[¹³CH₃]^δ² labeling of PDIA6 MLVpS: 240 mg of 2-hydroxy-2-[¹³C]methyl-3-oxo-4,4,4-tri-[²H]-butanoate (pro-S acetolactate-¹³C, NMR-Bio) and 100 mg of [¹³C]^ε-L-methionine (Sigma-Aldrich). For [U-²H, ¹⁵N, ¹²C], Met-[¹³CH₃]^ε, Val-[¹³CH₃]^γ2 labeling of PDIA6 MVpS: 240 mg of 2-hydroxy-2-[¹³C]methyl-3-oxo-4,4,4-tri-[²H]-butanoate (pro-S acetolactate-¹³C, NMR-Bio), 30 mg of L-leucine-d₁₀ and 100 mg of [¹³C]^ε-L-methionine (Sigma-Aldrich). For [U-²H, ¹⁵N, ¹²C], Met-[¹³CH₃]^ε, Val-[¹³CH₃/¹²C²H₃]^cγ1/cγ2, Ile-[¹³CH₃]^δ1 labeling of BiP IMV: 100 mg of 2-Keto-3-(methyl-d3)-butyric acid-4-¹³C,3-d sodium salt, 30 mg of L-leucine-d₁₀, 80 mg of α-ketobutyric acid methyl ¹³C (99%) 3,3-D2 (98%) and 100 mg of [¹³C]^ε-L-methionine (Sigma-Aldrich). One hour after the addition of the precursors, protein expression was induced by adding 0.5-1 mM IPTG at 25 °C for 12 hours. Methyl labeled proteins were purified following the protocols described above.

Mutagenesis, expression and purification of the PDIA6 and BiP mutants

The QuikChange II mutagenesis protocol (Stratagene) was used to introduce the PDIA6 mutations M182L, M331L, M359L, M367L, L61M, V75I, V124I, L139M, L165M, L196M, I234V, L320M, L345M, L349M, L385M, I406V, V407I, L419M, V421I, I425V, L427M, L432M, L435M, L440M, K161 to stop (domain a⁰), L139STOP (domain a⁰ Δlinker), P274 to stop (domain a⁰a, domain a), S428 to stop (domain b ΔC(13)), 420STOP (ΔC(20), domain b ΔC(20), domain ab ΔC(20)), E422QD423ND424ND426ND429NE431QD433ND434N (EQDN, dom b EQDN), R142QK150NR153Q, K150NR153QK159N and R142QK150NR153QK159NK160N (5x mutant) and the BiP mutation G407 to stop (NBD). Polymerase chain reaction primers were obtained from Microsynth. The plasmids were purified using the Zyppy plasmid mini prep kit from Zymo Research and mutations confirmed by sequencing from Microsynth AG. The expression and purification of the mutant proteins was performed as described for the wild-type proteins. All linker mutants (R142QK150NR153Q, K150NR153QK159N and R142QK150NR153QK159NK160N) behaved like the WT protein.

NMR spectroscopy

NMR experiments were performed in 25 mM Hepes (pH 7.5), 150 mM KCl, 10 mM MgCl₂ or 25 mM MES (pH 6.5), 150 mM KCl, 10 mM MgCl₂ at 37 °C. The experiments were recorded on Bruker Ascend 600 MHz, AscendII 700 MHz, Avance 800 MHz or Avance 900 MHz spectrometers running Topspin 3.6 and equipped with a cryogenically cooled triple-resonance probe. NMR data were processed with nmrPipe⁴⁴ and ccpnmr⁴⁵. Reference experiments were recorded at concentrations between 25 μM 100 μM. Secondary chemical shifts were calculated relative to random-coil values⁴⁶. Depending on the conditions, 10 mM TCEP, 10 mM CaCl₂, 5 mM ADP, 10 eq. linker peptide and 1 mM K₂HPO₄/KH₂PO₄ were used as indicated. To mimic ER-homeostatic like conditions 10 mM TCEP or 15 mM glutathione with GSSG:GSH 7:1 was used in presence of 10 mM Ca²⁺. Samples were incubated prior to the experiment (~1 h for TCEP, glutathione and ADP, ~30 min for protein-protein interaction experiments, Ca²⁺ and Gd³⁺, ON for peptides).

Assignment of PDIA6 Met[CH₃]^ε, Val[CH₃]^γ2 and Leu[CH₃]^δ² methyl groups

PDIA6 Met[CH₃]^ε, Val[CH₃]^γ2 and Leu[CH₃]^δ² assignment was obtained using a structure-based approach combining mutagenesis, specific valine labeling schemes⁴⁷, subconstructs and 3D ¹³C,¹³C-resolved [¹H,¹H]-NOESY experiments recorded at 37 °C in SEC buffer with 10 mM TCEP and 5% D₂O. The following point mutation were used: M182L, M331L, M359L, M376L, L61M, V75I, V124I, V139M, L165M, L196M, I234V, L320M, L345M, L349M, L385M, I406V, V407I, L419M, V421I, I425V, L427M, L432M, L435M, L440M. The following sub-constructs were used: his-domain a, his-domain ab. Specific valine labeling allowed to distinguish between leucine and valine residues. Each sample was recorded at 37 °C with an adjusted duration depending on the final concentration of each mutant (experimental time ranging from 120 to 240 min per sample). 3D ¹³C,¹³C-resolved [¹H,¹H]-NOESY experiments were recorded in 25 mM Hepes (pH 7.5), 150 mM KCl, 10 mM MgCl₂, 10 mM TCEP, 5% D₂O with sample concentrations of 870 μM with a mixing time of 500 ms. Together, this allowed for the assignment of 62/66 MetCε and LeuCδ2 and ValCγ2 methyl groups (94%).

Amide backbone assignment

For the sequence-specific backbone resonance assignments the following experiments were recorded: [U-¹H, ¹³C, ¹⁵N]-PDIA6 domain a⁰: HNCA and HNCACB at 37 °C in 25 mM MES (pH 6.5), 150 mM KCl, 10 mM MgCl2, 10 mM TCEP, 5 % D2O. [U-¹H, ¹³C, ¹⁵N]-PDIA6 his-domain a: 4D-APSY⁴⁸ at 37 °C in 25 mM MES (pH 6.5), 150 mM KCl, 10 mM MgCl2, 10 mM TCEP, 5 % D2O. [U-²H, ¹³C, ¹⁵N]-PDIA6 his-domain ab: HNCA, HNCACB, HNCO and HNcoCA at 37 °C in 25 mM MES (pH 6.5), 150 mM KCl, 10 mM MgCl2, 10 mM TCEP, 5 % D2O and HNCA, HNCACB with deuterium decoupling and 3D ¹⁵N, ¹⁵N resolved [¹H, ¹H]-NOESY with a mixing time of 100 ms at 37 °C in 25 mM Hepes (pH 7.5), 150 mM KCl, 10 mM MgCl2, 10 mM TCEP, 5 % D2O. [U-¹H, ¹³C, ¹⁵N]-PDIA6 his-domain bΔC: HNCA, HNCACB, HNcoCACB and HNCO at 25 °C in 25 mM HEPES (pH 7.5), 150 mM KCl, 10 mM MgCl2, 10 mM GSSG, 5 % D2O and HNCACB, HNcaCO and 3D ¹⁵N, ¹⁵N resolved [¹H, ¹H]-NOESY with a mixing time of 80 ms at 25 °C in 25 mM HEPES (pH 7.5), 150 mM KCl, 10 mM MgCl2, 10 mM TCEP, 5 % D2O. All experiments were recorded at sample concentrations of 0.5-1 mM. The amide backbone assignments comprise 85% of domain a⁰, 92% of his-domain a, 79% of his-domain b and 54% of his-domain ab.

Fluorescent labeling of purified recombinant PDIA6 and BiP NBD

All PDIA6 constructs were N-terminally tagged with DyLight^TM 488 Amine-Reactive Dyes (Thermo Scientific) at a concentration of 10 mg/mL in 25 mM Mes (pH 6.5) 150 mM KCl, 10 mM MgCl₂ in the dark for 1 h at 25 °C. Free dye was removed by dialysis at pH 6.5 at 4 °C in the dark overnight and the labelled proteins buffer exchanged (3x) to SEC buffer. BiP NBD was N-terminally tagged with DyLight^TM 633 Amine-Reactive Dyes (Thermo Scientific) following the same protocol.

SEC-MALS

SEC-MALS measurements of PDIA6 constructs were performed at 25 °C in SEC buffer (25 mM Hepes, pH 7.5, 150 mM KCl and 10 mM MgCl₂) using a S200 10/300 GL column (Agilent Technologies) on an Agilent 1260 HPLC. Proteins eluted at a concentration of 20 μM. Elution was monitored by multiangle light scattering (Heleos II 8+; Wyatt Technology), differential refractive index (Optilab T-rEX; Wyatt Technology) and absorbance at 280 and 254 nm (1260 UV; Agilent Technologies). The column was equilibrated overnight in the running buffer to obtain stable baseline signals from the detectors before data collection. All the system parameters were calibrated using an injection of 2 mg/ml BSA solution (ThermoPierce) and standard protocols in ASTRA 6. Molar mass and mass distributions were calculated using the ASTRA 6 software (Wyatt Technology).

Differential scanning fluorimetry

DSF data were acquired using a Prometheus NanoTemper (Prometheus NT.48 instrument, NanoTemper Technology) at protein concentrations of 100 μM in SEC buffer. The samples were scanned from 20 °C to 95 °C at a scan rate of 0.5 °C/min. Protein unfolding was measured by detecting the temperature-dependent change in tryptophan fluorescence at emission wavelengths of 330 nm and 350 nm. Melting temperatures were determined by detecting the maximum of the first derivative of the fluorescence ratios (F350/F330).

Microscale thermophoresis experiments

For the determination of the dissociation constants N-terminally tagged DyLight 488 PDIA6 at 20 nM was incubated with increasing amounts of BiP in SEC buffer in presence of 5 mM ADP, 1 mM KH₂PO₄/K₂HPO₄, 0.05% Tween and 1 mM CaCl₂ prepared by serial dilution. Samples were incubated for 30 min at 25 °C prior to loading into capillary tubes. The LED power was set to 100% and the MST power to medium. The pre-MST period was 5 s, the MST-acquisition period was 30 s, and the post-MST period was 5 s. Measurements were performed in triplicates at a Monolith NT.115 (NanoTemper) and integrated at a 2.5 s interval.

Mass photometry

Mass photometry data were obtained using a Refeyn OneMP (Refeyn Ltd.) in microscope coverslips prepared according to the manufacturer’s instructions with silicone gaskets. 18 µl of SEC buffer were used to lock the focus, then 2 µl protein sample at indicated final concentrations were added. Movies were recorded using Refeyn Acquire^MP 2.5.1 and analysed with Refeyn Discover^MP 2.5.0.

Crystallization, data collection and structure determination of PDIA6 domain bΔ

PDIA6 his-domain bΔC (P274-L427) in 25 mM Hepes (pH 7.5), 150 mM KCl, 5 mM MgCl₂, 10 mM TCEP was crystallized with 10 mg/mL at room temperature in sitting-drop vapor diffusion experiments in a 1:1 ratio of protein and precipitant. Crystals appeared in 1.9 M (NH4)₂SO₄, 0.1 M Hepes 7.0, 1% DMSO after 2 days and grew to their final size within 10 days. Crystals were cryo-preserved by addition of ethylene glycol to a final concentration of 20% (v/v) and flash cooled in liquid nitrogen. Data was collected at the SLS beamline X06DA (Swiss Light Source, Paul Scherrer Institute, Switzerland) at 100 K and were integrated, indexed and scaled using XDS software^49,50. The crystal structure was determined by molecular replacement with Phaser⁵¹ using PDB: 3uem⁵². Automated model building⁵³ was carried out with Arp/Warp⁵⁴ followed by Autobuild in PHENIX⁵⁵, and was completed by manual model building with Coot⁵⁶ and structure refinement in PHENIX⁵⁷. Model quality was validated with Molprobity⁵⁸. Data collection and refinement statistics are summarized in Table S2. The atomic coordinates and structure factors have been deposited in the Protein Data Bank under the accession code 8cpq.

In vitro liquid-liquid phase separation

PDIA6 LLPS was examined at 25 μM in SEC buffer in presence of 10% polyethylene glycol (PEG3350, mass fraction, freshly prepared directly prior to experiment) as a crowding agent in absence and presence of 15 mM glutathione with GSSG:GSH 7:1 or 10 mM TCEP, 10 mM CaCl₂ and/or 20 mM EDTA. PEG was added last by simultaneously swirling the sample and gently pipetting up and down to thoroughly mix the sample. Images were taken 30 min after addition of PEG. No PDIA6 construct formed liquid droplets in the absence of crowding agent and at concentrations below 1 mM. Phase separated PDIA6 droplets formed under ER-mimicking conditions (15 mM glutathione with GSSG:GSH 7:1 + 1-10 mM CaCl₂) as well as under strongly reducing conditions (10 mM TCEP) in presence of 1-10 mM CaCl₂. Several salt concentrations (50-350 mM) were tested. Unlabeled proteins were used to confirm that they undergo phase separation without fluorescent labelling. The 96 well glass bottom plate were coated with 1% Pluronic F-127 overnight and the wells washed three times with SEC buffer. For fluorescence microscopy 1% N-terminally tagged DyLight protein was added to the unlabeled protein. BiP NBD LLPS was examined at 20 μM in SEC buffer in presence of 5 mM ADP and 10% polyethylene glycol as a crowding agent in absence and presence of 20 μM PDIA6. Samples including BiP NBD were prepared as described for PDIA6.

In vitro fluorescence microscopy

A widefield microscope FEI MORE with a Hamamatsu ORCA flash 4.0 cooled sCMOS camera using a TIRF APON 60x/1.49 and U Plan S Apo 100x/1.4 oil objective with Live Acquisition 2.5 software was used for capturing images. Pictures were processed in OMERO. All images are representative from at least three independent sets of experiments. Post-acquisition coloring of the channels according to artistic preference.

Plasmid generation for cell transfection

To assure ER-localization, the KDEL signal sequence was cloned at the C-terminus of PDIA6-tGFP (Origene #RG201710) using NEBuilder HiFi Assembly cloning kit (New England Biolabs, NEB) with primers designed by the NEBuilder Assembly Tool following manufacturer’s instructions, for simplicity, we refer to this plasmid as PDIA6-GFP. The C-terminal tail of PDIA6 was truncated by removing the last twelve amino acids from its sequence in the PDIA6-tGFP-KDEL plasmid using NEB site-directed mutagenesis kit (NEB) following manufacturer’s instructions and primers selected using NEBaseChanger tool. The PDIA6-EGFP-KDEL plasmid for transient transfections was generated from PDIA6-tGFP-KDEL plasmid and the EGFP insert from ACE2-EGFP (Addgene #154962), using NEBuilder HiFi Assembly cloning kit (New England Biolabs, NEB) with the primers designed by the NEBuilder Assembly Tool following manufacturer’s instructions. The QuikChange II mutagenesis protocol (Stratagene) was used to introduce the mutations K159NK160N and R124QK159NK160N. Mutagenesis of L128AL131AL134AV135AL139A, R142QK150NR153QK and R142QK150NR153QK159NK160N was performed by GenScript. PDIA6 L128AL131AL134AV135AL139A has been characterized to be monomeric²³ and is hence referred to as “PDIA6 monomer” in the manuscript. Human preproinsulin followed with a C-terminal myc-His A tag was synthesis by BioCat. The genes were inserted using BamHI and XhoI restriction enzymes into a pcDNA3.1(+) expression vector (Novagen). The QuikChange II mutagenesis protocol (Stratagene) was used to introduce the Akita proinsulin mutation C96Y. Hamster BiP-mCherry was a gift from Erik Snapp (Addgene plasmid #62233) and mutated to the human sequence. All polymerase chain reaction primers were obtained from Microsynth. The plasmids were purified using the Zyppy plasmid mini prep kit from Zymo Research and mutations confirmed by sequencing from Microsynth AG.

Cell culture

HeLa CCL2, HEK293A and U2OS cells were grown at 37 °C and 5% CO₂ in high-glucose Dulbecco’s modified Eagle’s medium (DMEM, Sigma-Aldrich) supplemented with 10% fetal bovine serum (FBS, Biowest), 2 mM L-Glutamine (Gibco), 1 mM Sodium Pyruvate (Sigma), and 1 x Penicillin and Streptomycin (Sigma). HeLa CCL2 and HEK293A cell lines were a kind gift of Prof. Dr. Martin Spiess, with their identities authenticated by STR analysis by Microsynth AG (Balgach, Switzerland). U2OS cell lines were a kind gift of Prof. Dr. Michael Hall. All cell lines were confirmed to be mycoplasma-negative by PCR. For transient cell transfections, cells were plated into 6-well plates to reach 70% confluency the following day and transfected with 1 µg plasmid DNA complexed with Helix-IN transfection reagent (OZ Biosciences). Cells were replated onto coverslips the following day and fixed 24-48 h post transfection. For stress treatments, cells were treated with tunicamycin (1 μM), thapsigargin (1.25 μg/mL) or cyclopiazonic acid (8 μM) for specified durations. For washout experiments, cells were treated for 10 h with tunicamycin (1 μM), thapsigargin (1.25 μg/mL) or cyclopiazonic acid (8 μM), washed with PBS and cultured in fresh medium overnight.

Immunostaining

HeLa cells were plated onto coverslips 24 hours prior to fixation or stress treatment. At specified time points, cells were fixed in 4% paraformaldehyde, permeabilized with 0.1 % Triton X-100, blocked in PBS containing 5%FBS, and stained with anti-PDIA6 (1:200, GeneTex GTX33397), anti-GRP78/BiP (1:200, Invitrogen 1H11-1H7), anti-calreticulin (1:200, Proteintech 27298-1-AP), anti-DNAJB11/ERdj3 (1:200, Thermofisher #15484-1-AP), anti-Grp94 (1:200, Enzo SPA-850), anti-PDIA1 (1:200, Genetex GTX2279), anti-calnexin (1:200, Enzo) followed by AF488-conjugated or AF633-conjugated goat anti-rabbit, AF633-conjugated goat anti-mouse, AF488-conjugated goat anti-rabbit or AF633-conjugated goat anti-rabbit, AF488-conjugated goat anti-rabbit or AF633-conjugated goat anti-rabbit, AF488-conjugated goat anti-rat or AF546-conjugated donkey anti-rat, AF633-conjugated goat anti-mouse and AF488-conjugated or AF633-conjugated goat anti-rabbit, respectively. Proinsulin was visualized within the condensates by staining preproinsulin-myc-His transfected HeLa cells with anti-myc clone 9E10 (1:2000, Sigma 9E10) primary antibody followed by AF633-conjugated anti-mouse. Overexpressed PDIA6-GFP was imaged in the 488 channel using the eGFP signal. Coverslips were mounted onto glass slides with Fluoromount G (Southern Biotech) and sealed with nail polish.

Microscopy of fixed cells

Confocal images were acquired with Olympus Fluoview FV3000 system, using an UPLSAPO 60x/1.30 objective with silicone oil using the FV3000 (FV21S-SW Version 2.5.1) system software. Laser intensities were at 0.5–3% for both 488 (AF488) and 640 (AF633) wavelengths. Sampling speed was 8.0 μs/pixel. All images are representative from at least three independent sets of experiments. All images for corresponding experiments were processed with the same settings to insure comparable results. Post-acquisition coloring of the channels according to artistic preference.

Live-cell imaging

For live-cell imaging, cells were seeded on imaging chamber (ibidi μ-slide) 24 h prior to data acquisition. Live-cell imaging was performed in complete growth medium lacking phenol red at 37 °C with 5% CO₂ on a widefield microscope FEI MORE with a Hamamatsu ORCA flash 4.0 cooled sCMOS camera using a TIRF APON 60x/1.49 oil objective and a Olympus „SpinD“ spinning disc confocal microscope with a Hamamatsu ORCA-Fusion, sCMOS using an UPL APO 60x/1.5 oil objective. Laser intensities were set to 5-10% for 488 wavelength. The system was preheated overnight (MORE) or 1 h (SpinD) to 37 °C to prevent thermal fluctuations. All images are representative from at least three independent sets of experiments. Images of the movies were recorded every 0.5-1.0 s. Post-acquisition coloring of the channels according to artistic preference.

FRAP

FRAP experiments were performed on a Olympus „SpinD“ spinning disc confocal microscope with a Hamamatsu ORCA-Fusion, sCMOS using an UPL APO 60x/1.5 oil objective. Droplets and condensates were bleached using a 488 nm laser with 100% intensity (Rapp OptoElectronic Firefly). A total 10 images were obtained prebleaching, while 200 images were recorded every 1.0 -1.83 s (in vitro) or 0.5-1.0 s (in vivo) postbleaching. Intensities of FRAP regions were extracted with Fiji 1.52p software tool, background corrected and reported relative to the prebleaching time point.

Image analysis

Pearson Correlation coefficients were determined using the Fiji “JaCOP” plugin from 10-15 cells (see Table S2 for details). Statistical significance was calculated using the two-tailed Student’s t-test. The fluoresce intensity histograms were extracted using the Fiji 1.52p software “plot profile” tool.

Western blotting and antibodies

To assess PDIA6 and BiP expression levels, HeLa CCL2 cells were lysed in lysis buffer (1% Triton X-100, 150 mM NaCl, 20 mM Tris pH 7.5, 1mM EDTA, 1mM EGTA, protease inhibitor) and denatured in Laemmli buffer at 65 °C for 10 min. The three biological replicates were prepared and lysed independently (n=3). Samples were resolved by 10% SDS-PAGE and transferred onto nitrocellulose membrane (Amersham). Membranes were blocked with TBST (20 mM Tris, 150 mM NaCl, pH 7.6, 0.1% Tween20) with 5% non-fat dry milk for 30 min and incubated with anti-PDIA6 primary antibody (1:2000, GeneTex GTX33397), anti-GRP78/BiP primary antibody (1:2000, Invitrogen 1H11-1H7) or anti-tubulin primary antibody (1:2000, Sigma T5168) overnight at 4°C, followed by 2 h incubation with HRP-conjugated secondary antibody (1:20000; anti-mouse or anti-rabbit, Invitrogen 31430 and 31460) in TBST. Chemiluminescence signals were detected using Immobilon Western HRP Substrate (Advansta) and imaged using a FusionFX (Vilber Lourmat).

To asses insulin secretion, cells were transfected with pre-proinsulin (and PDIA6 constructs). After 24 h cells were washed with preheated PBS (1x) and OPTIMEM Glutamax (Gibco) (4x) and incubated in OPTIMEM Glutamax (16 h). The medium was collected, spun down (10 min, 13’000 rpm, 4°C) and the supernatant precipitated by the addition of 100% TCA (1 volume TCA to 4 volume protein). After 10 min incubation on ice, the samples were spun down (5 min, 13’000 rpm, 4°C) and washed twice with cold acetone. The dried pellet was dissolved in 30 μL of Laemmli buffer by vortexing and boiling (10 min, 300 rpm, 95°C). The cells were lysed in RIPA buffer and denatured in Laemmli buffer at 65 °C for 10 min. The four biological replicates were prepared and lysed independently (n=4). Protein levels were measured using a BCA assay (Merck) and the volumes of the precipitate and lysates adjusted to equal lysate protein concentrations. Samples were resolved by 4-20% SDS-PAGE (Biorad) and transferred onto nitrocellulose membranes (Biorad). The lysate blot was processed as described above. The blot of the secreted proteins was preblocked with TBST with 5% non-fat dry milk for 5 min and the samples fixed by incubation with TBST with 0.2% glutaraldehyde for 15 min. The membrane was washed in TBST, blocked with TBST with 5% non-fat dry milk for 30 min and incubated with anti-pro/insulin antibody (1:1000, Invitrogen MA1-83256) in Can get signal (Toyobo) overnight at 4°C, followed by 2 h incubation with HRP-conjugated secondary antibody (1:20000; anti-mouse, Invitrogen 31430) in Can get signal solution. Chemiluminescence signals were detected using Immobilon Western HRP Substrate (Advansta) and imaged using a FusionFX (Vilber Lourmat).

MTT assay

HeLa cells were seeded in 96-well plate 24 h after transfection. After 24 h cells were stressed with Tm and MTT (Merck) was added at indicated timepoints at a final concentration of 500 μg/mL and incubated for 2 h at 37 °C. The medium was removed and the crystals dissolved in acidic (0.08 M HCl) isopropanol. The absorbance (A_550mm- A_690mm) was measured on a plate reader. All experiments were repeated in experimental and independent biological triplicates. Significance was calculated using the two-tailed Student’s t-test (n=9).

RT-qPCR

RNA was extracted and purified from cells using RNeasy kit following manufacturer’s instructions. cDNA was reverse-transcribed using GoScript reverse transcriptase primed with a mix of Oligo(dT)s and random hexamers (Promega). RT-qPCR was performed using GoTaq qPCR master mix (Promega) and primers specific for sXBP1. sXBP1 expression was normalized for β-actin expression. For PDIA6 WT transfected cells, the sXBP1 levels were still below detection limit after 40 cycles of qPCR and Ct was thus set to 40 for statistical analysis. The significance was calculated from three independent biological replicates using the two-tailed Student’s t-test (n=3).

Structural models

BiP structural model in the ADP undocked states was obtained by homology modelling using the SWISS-MODEL server using as a template the DnaK undocked conformation (PDB 2kho)⁵⁹. The structural model of PDIA6 FL has been modeled as a chimera of the individual domain crystal structures (domain a⁰ (PDB: 4ef0), domain a (PDB: 4gwr), domain b (PDB: 8cpq, determined in this study)). The linker connecting domain a⁰ and a and C-terminal tail were modeled as disordered based on the NMR data presented in Fig. 2 and Extended Data Fig. 3-4. The interface between the domain a⁰ monomers is based on the crystal structure of domain a⁰ (PDB: 4ef0). The interface between domains a and b was modeled using ESMFold⁶⁰ based on the NOE contact between the domains presented in Fig. 2. The binding interface of the linker to domain b was modeled based on the experiments presented in this manuscript.

Structural comparison of helix α4

The following protein structures have been used to compare the length of helix α4: PDB: 4ekz (PDIA1 a and a’), 2dmm (PDIA3 a’), 2dj1 (PDIA4 a⁰), 2dj2 (PDIA4 a), 2dj3 (PDIA4 a’), 4ef0 (PDIA6 a⁰) and 4gwr (PDIA6 a).

Data availability

All data needed to evaluate the conclusions of the paper are presented in the manuscript. PDIA6 MLVpS methyl and his-domain a⁰, his-domain a and his-domain ab amide backbone resonance assignments have been submitted to the Biological Magnetic Resonance Data Bank under the following accession codes: 51863 and 51864. The crystal structure of the PDIA6 domain b has been deposited in the protein data bank under the accession code: 8cpq. Additional data related to this paper will be provided by the authors upon request.

Acknowledgements

We thank Timothy Sharpe, Tobias Mühlethaler, Kai Schleicher, Sara Roig and Thomas Müntener for technical support and Bruno Reversade and Elif Karagöz for fruitful discussions.

Author Contributions

A.L., G.M., A.S. and S.H. designed the study. G.M., A.S., T.M. and S.H. supervised the work. A.L. expressed and purified PDIA6 constructs and performed biochemical, biophysical and LLPS characterizations. A.L and G.M. expressed and purified BiP protein. A.L. and G.M. recorded and analyzed NMR data. A.L. performed the in cell experiments. A.L., G.M., V.S. and A.S. analyzed and discussed the cell biology data. R.P.J. solved the crystal structure. A.L., G.M. and S.H. wrote the manuscript with contributions from all authors. This work was funded by the Swiss National Science Foundation (grant 207755 to S. H.).

Competing interests

The authors declare no competing interests.

Materials and Correspondence

Correspondence to [email protected] and [email protected]

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A functional chaperone condensate in the endoplasmic reticulum

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