A structured interdomain linker directs self-polymerization of human uromodulin
Edited by Paul Wassarman, Mount Sinai School of Medicine, New York, NY, and accepted by the Editorial Board December 23, 2015 (received for review October 6, 2015)
Significance
Urinary tract infection is the most common nonepidemic bacterial infection in humans, with 150 million cases per year and a global health care cost above $6 billion. Because the urinary tract is not protected by mucus, mammals produce a molecular net that captures pathogenic bacteria in the urine and clears them from the body. By visualizing the 3D structure of its building block, glycoprotein uromodulin, we provide insights into how the net is built, and how it is compromised by mutations in patients with kidney diseases. Our work also explains nonsyndromic deafness due to mutations affecting the tectorial membrane, a similar filamentous structure in the human inner ear.
Abstract
Uromodulin (UMOD)/Tamm–Horsfall protein, the most abundant human urinary protein, plays a key role in chronic kidney diseases and is a promising therapeutic target for hypertension. Via its bipartite zona pellucida module (ZP-N/ZP-C), UMOD forms extracellular filaments that regulate kidney electrolyte balance and innate immunity, as well as protect against renal stones. Moreover, salt-dependent aggregation of UMOD filaments in the urine generates a soluble molecular net that captures uropathogenic bacteria and facilitates their clearance. Despite the functional importance of its homopolymers, no structural information is available on UMOD and how it self-assembles into filaments. Here, we report the crystal structures of polymerization regions of human UMOD and mouse ZP2, an essential sperm receptor protein that is structurally related to UMOD but forms heteropolymers. The structure of UMOD reveals that an extensive hydrophobic interface mediates ZP-N domain homodimerization. This arrangement is required for filament formation and is directed by an ordered ZP-N/ZP-C linker that is not observed in ZP2 but is conserved in the sequence of deafness/Crohn’s disease-associated homopolymeric glycoproteins α-tectorin (TECTA) and glycoprotein 2 (GP2). Our data provide an example of how interdomain linker plasticity can modulate the function of structurally similar multidomain proteins. Moreover, the architecture of UMOD rationalizes numerous pathogenic mutations in both UMOD and TECTA genes.
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Uromodulin (UMOD) is expressed in the thick ascending limb of Henle’s loop as a GPI membrane-anchored precursor that consists of three EGF-like domains, a domain of unknown function (D8C), and a zona pellucida (ZP) module (1, 2) (Fig. 1A, Top). The latter, containing Ig-like domains ZP-N and ZP-C (3–5), is found in other medically important human glycoproteins linked to infertility (egg coat components ZP1–ZP4), nonsyndromic deafness [inner ear α- and β-tectorin (TECTA/B)], Crohn’s disease [glycoprotein 2 (GP2)], and cancer [TGF-β coreceptors betaglycan (BG) and endoglin (ENG)] (6, 7). Upon processing by Ser protease hepsin (8) at a consensus cleavage site (CCS) C-terminal to the ZP module (9), UMOD sheds a C-terminal propeptide (CTP) that contains a polymerization-blocking external hydrophobic patch (EHP), exposing an internal hydrophobic patch (IHP). This event triggers homopolymerization into filaments that are excreted into the urine (4, 10), where UMOD performs a plethora of biological functions, including protection against urinary tract infections, prevention of kidney stones, and activation of innate immunity (1, 2, 11, 12).
Fig. 1.
Although UMOD activity is strictly linked to its supramolecular state (2), the mechanism of ZP module-dependent assembly remains unclear. Mass spectroscopy (MS) analysis of ZP-C disulfide linkages suggests that there are two types of ZP modules with different structures (13). Type II contains 10 conserved Cys (C1–7,a,b,8) and both homopolymerizes (UMOD, GP2, and TECTA) and heteropolymerizes (ZP1, ZP2, and ZP4), whereas type I (ZP3) includes eight conserved Cys (C1–8) and only heteropolymerizes with type II (7, 13, 14). However, MS studies of egg coat protein disulfides are contradictory (15), and type II disulfide linkages C5–C6, C7–Ca, and Cb–C8 are compatible neither with the fold of ZP3 (3) nor with structures of the ZP-C domain of BG, whose ZP module contains 10 Cys (16, 17). At the same time, interpretation of the latter data in relation to polymerization is complicated by the fact that, like ENG, BG remains membrane-associated and does not form filaments (7, 17).
To gain insights into the mechanism of ZP module protein assembly, we carried out X-ray crystallographic studies of the complete polymerization region of UMOD. The structure reveals that a rigid interdomain linker is responsible for maintaining UMOD in a polymerization-competent conformation. This rigid linker is conserved in homopolymeric ZP modules, but it is flexible in the structure of ZP2, also presented in this work, which, together with ZP3, forms heteropolymeric egg coat filaments. Furthermore, ZP module proteins that do not make filaments lack such a linker. Because UMOD and ZP2 show conservation of both disulfide pattern and fold, our data reveal that the interdomain linker, rather than a different ZP-C structure, underlies the ability of UMOD to self-assemble. Accordingly, polymerization-competent UMOD forms a dimer via β-sheet extension and hydrophobic interactions, and disruption of this dimer interface completely abolishes filament formation. Our study yields insights into the formation of an essential polymerization intermediate of UMOD and highlights how an interdomain linker can regulate the biological function of a multidomain protein.
Results and Discussion
Maltose-Binding Protein-Fused UMOD Secreted by Mammalian Cells Polymerizes Like Native UMOD.
To shed light on UMOD polymerization, we focused on a protease-resistant fragment (residues S292–F587) that contains the ZP module (Fig. 1A, Top), constitutes the core of UMOD filaments (18), and matches an alternatively spliced isoform of GP2 (19). UMODp (residues S292–Q640), a related construct that includes the C-terminal GPI-anchoring site, was expressed in mammalian cells as a fusion with a mammalianized version of bacterial maltose-binding protein (mMBP) (Fig. 1A, Middle). Electron microscopy (EM) revealed that secreted mMBP-UMODp forms native-like filaments with the characteristic zigzag structure of urinary UMOD (20), full-length recombinant UMOD, or elastase-treated UMOD (Fig. 1 B–E).
Crystal Structure of the Polymerization Region of UMOD.
Despite extensive attempts, we could not obtain diffracting crystals of depolymerized native UMOD or unfused recombinant UMOD constructs. However, a soluble version of mMBP-UMODp (including UMOD residues G295–Q610) that cannot be cleaved at the CCS and carries a mutation of nonessential glycosylation site N513 (mMBP-UMODpXR; Fig. 1A, Bottom) formed crystals in high-salt conditions (Fig. S1A). The structure of mMBP-UMODpXR, with two molecules per asymmetrical unit, was solved by molecular replacement with MBP as a search model and refined to R = 22.1%, Rfree = 24.6% at a resolution of 3.2 Å (Fig. 2A and Table S1). The entire molecule A has well-defined electron density (Fig. S1B), which reveals that a fourth EGF-like domain precedes the ZP module of UMOD (Fig. 2A). This domain is structurally most similar to human TGF-α (21), with a root-mean-square deviation (rmsd) of 1.4 Å over 23 residues. Not visible in molecule B due to flexibility within the crystals rather than proteolytic degradation (Fig. S2), EGF IV consists of a short N-terminal α-helix and an antiparallel β-turn disulfide bonded with C1′–C3′ and C2′–C4′ connectivity (Fig. 2B). Mutations of the corresponding Cys are associated with autosomal dominant tubulointerstitial kidney disease (ADTKD) (Fig. S3 and Table S2). An additional C5′–C6′ disulfide tethers EGF IV C317 to ZP-N C347, which belongs to an α-helix that lies between strands B and C (Fig. 2B) and is absent in ZP3 (3, 5). Loss of either Cys is also associated with ADTKD, due to intracellular aggregation and impaired urinary secretion of UMOD (22, 23) (Fig. S3 and Table S2). Interestingly, human GP2 and TECTA, as well as chicken ZPD [a peripherally associated homopolymeric egg coat component (24)], also contain an EGF IV-like Cys-rich domain N-terminal to their ZP module (Fig. S3). Taken together, these data identify a subset of sequence-related but functionally diverse proteins that are characterized by EGF and ZP-N domains linked by a disulfide bond.
Fig. 2.
Fig. S1.
Table S1.
Crystal (PDB ID code) | mMBP-UMODpXR (4WRN) | ZP2 ZP-C (5BUP) |
---|---|---|
Experiment | ||
Beamline | ESRF ID29 | DLS I02 |
Wavelength, Å | 0.97625 | 0.97939 |
No. of crystals | 1 | 1 |
Data collection | ||
Space group | H32/155 | P61/169 |
Cell dimensions | ||
a, b, c; Å | 242.32, 242.32, 258.86 | 105.21, 105.21, 40.55 |
α, β, γ; ° | 90, 90, 120 | 90, 90, 120 |
Molecules, A.U. | 2 | 1 |
Solvent content, % | 73.7 | 54.5 |
Mosaicity, ° | 0.104 | 0.183 |
Wilson B factor, Å2 | 108.3 | 33.3 |
Resolution, Å | 50.0–3.20 (3.28–3.20) | 45.56–2.25 (2.31–2.25) |
Total reflections | 300,327 (20,429) | 41,295 (2,855) |
Unique reflections | 47,986 (3,370) | 12,296 (866) |
Completeness, % | 99.0 (94.5) | 99.4 (99.7) |
Redundancy | 6.3 (6.1) | 3.4 (3.3) |
I/σI | 16.20 (1.16) | 9.75 (1.13) |
CC(1/2), % | 99.9 (43.2) | 99.6 (51.1) |
Rpim, % | 3.8 (65.5) | 6.5 (67.7) |
Refinement | ||
Resolution, Å | 34.98–3.20 (3.31–3.20) | 45.56–2.25 (2.37–2.25) |
Reflections | 47,792 (4,720) | 11,813 (1,536) |
Free reflections | 2,431 (264) | 1,174 (149) |
Rwork/Rfree, % | 22.10 (40.12)/24.59 (40.37) | 20.14 (34.30)/22.83 (36.25) |
CCwork/CCfree | 0.938 (0.632)/0.915 (0.563) | 0.955 (0.617)*/0.934 (0.672)* |
ML coordinate error, Å | 0.49 | 0.33 |
ML phase error, ° | 28.21 | 26.02 |
rmsd | ||
Bond lengths, Å | 0.004 | 0.005 |
Bond angles, ° | 0.840 | 0.859 |
Ramachandran plot | ||
Favored, % | 99.0 | 98.0 |
Allowed, % | 1.0 | 2.0 |
Outlier, % | 0.0 | 0.0 |
No. of atoms | ||
Total | 10,573 | 1,349 |
Protein | 10,483 | 1,266 |
Ligand/ion | 90 | 12 |
Water | 0 | 71 |
Protein residues | 1,356 | 159 |
Average B factor, Å2 | ||
Total | 141.3 | 46.9 |
Protein | 141.2 | 47.0 |
Ligand/ion | 152.1 | 81.8 |
Water | — | 39.0 |
Parameters for the outermost shell are shown in parentheses. A.U., asymmetric unit; CC(1/2), percentage of correlation between intensities from random half-datasets; CCfree, correlation of the experimental intensities of free reflections excluded from the refinement with the intensities calculated from the refined molecular model; CCwork, correlation of the experimental intensities with the intensities calculated from the refined molecular model; DLS, Diamond Light Source; ESRF, European Synchrotron Radiation Facility; I/σI, signal-to-noise ratio; ML, maximum likelihood; Rpim, Σhkl √(1/n − 1) Σi|Ii(hkl) − I(hkl)|/ΣhklΣi Ii(hkl), where Ii(hkl) is the intensity for an observation of a reflection and I(hkl) is the average intensity of all symmetry-related observations of a reflection.; Rwork, Σhkl‖Fobs| − k|Fcalc‖/Σhkl|Fobs|; Rfree, same as Rwork calculated from free reflections excluded from refinement.
*
Outer shell: 2.33–2.25 Å.
Fig. S2.
Fig. S3.
Table S2.
Protein | Mutation | Predicted effect |
---|---|---|
UMOD | C297W/Y | Destroys conserved disulfide bond C1′–C3′ |
C300G/Y/R | Destroys conserved disulfide bond C2′–C4′ | |
C306Y | Destroys conserved disulfide bond C1′–C3′ | |
K307T | Disrupts interaction with Q319 and Q316 | |
C315R/Y | Destroys conserved disulfide bond C2′–C4′ | |
R312C | Interferes with EGF IV disulfide bond formation | |
Q316P | Disrupts interaction with K307 and Q316 | |
C317Y | Destroys conserved disulfide bond C5′–C6′ between ZP-N and EGF IV, and thereby the coordination between the two domains | |
C347G | Destroys conserved disulfide bond C5′–C6′ between ZP-N and EGF IV, and thereby the coordination between the two domains | |
E375Q | Disrupts salt bridge with E304, and thereby the orientation between ZP-N and EGF IV | |
V458L | Bulky residue in the hydrophobic core of ZP-C interrupts hydrophobic sheet stacking | |
A461E | Bulky residue disrupts packing of linker α1 against the hydrophobic sheet of ZP-C, affecting ZP-N/ZP-C domain orientation | |
T469M | Disrupts interaction with T465, and thereby orientation between IHP and βA′ | |
G488R | Bulky residue disrupts packing of linker α1 against the hydrophobic sheet of ZP-C, affecting ZP-N/ZP-C domain orientation | |
T605G | Disrupts EHP conformation due to loss of interaction with V477 backbone | |
TECTA | P1791R | Disrupts the linker between EGF and ZP-N domains, thereby affecting their relative orientation |
L1820F | Disrupts αBC that harbors the EGF/ZP-N disulfide | |
G1824D | Disrupts the linker of αBC, harboring the EGF/ZP-N disulfide | |
C1837G/R | Destroys conserved disulfide bond C2–C3 | |
T1866M | Removes the highly conserved N-linked glycan site involved in dimerization | |
H1867R | Disrupts interaction with conserved T1866 and N-glycan, thereby interfering with dimer formation | |
Y1870C | Interferes with ZP-N C1–C4 disulfide bond | |
T1873I | Destroys interaction with E1895 | |
R1890C | Odd Cys interferes with disulfide bond formation and correct folding | |
C1898R | Destroys conserved disulfide bond C1–C4 | |
R1947C | Odd Cys interferes with disulfide bond formation and correct folding | |
A1982D | Charged residue interferes with hydrophobic sheet-sheet packing in the ZP-C core | |
I1997T | Polar residue interferes with hydrophobic sheet-sheet packing in the ZP-C core | |
D2006Y | Bulky residue in the conserved βC′′–βD loop disrupts interactions with and conformation of the loop/disulfide N-terminal to the CTP | |
I2009T | Polar residue interferes with hydrophobic sheet-sheet packing in the ZP-C core | |
R2021H | Disrupts the interaction with conserved E2013 and interface between ZP-N, ZP-C, and ZP-N/ZP-C linker affecting ZP-N/ZP-C domain orientation |
The ZP-N domain of UMOD (Figs. S1C and S4A) is similar to the ZP-N domain of ZP3 (Fig. S4 B and C), including invariant disulfides (5) and a conserved Tyr (Fig. S4, arrow) whose mutation in TECTA is associated with hearing loss (25). Moreover, it contains an N-linked glycosylation site (N396; Fig. 2A) that is also found in GP2 and TECTA (Fig. S3) as well as additional ZP module proteins, including ZPD (26, 27) (Fig. S3), olfactorin (28), pirica (29), larval glycoprotein (30), and SPP120 (31).
Fig. S4.
Surprisingly, our crystallographic data reveal that UMOD ZP-C (Figs. S1D and S5A) also shares the same fold and disulfide connectivity of ZP3 and BG ZP-Cs (3, 16, 17) (Fig. S5 B and C), except for the Ca–Cb disulfide not found in ZP3 proteins (15) (Fig. S5D). Accordingly, analysis of ZP-C Cys covariation based on multiple sequence alignments in Pfam (32) is consistent with C5–C7, C6–C8, and Ca–Cb connectivity (Fig. S5E).
Fig. S5.
The Crystal Structure of ZP2 ZP-C Reveals That ZP Modules Have a Conserved Disulfide Connectivity.
To confirm the existence of a single ZP module disulfide connectivity, we determined a 2.25-Å resolution structure of the ZP-C domain of mouse ZP2 (residues D463–D664; Fig. 3A and Fig. S6A). This molecule, which plays a key role in mammalian gamete recognition (33), has so far eluded structural determination but was reported to contain the alternative pattern based on C7–Ca, Cb–C8 MS assignments (13). The structure (R = 20.1%, Rfree = 22.8%; Fig. 3B, Fig. S6B, and Table S1) conclusively shows that ZP2 adopts the same disulfide linkages and overall fold as UMOD, ZP3, and BG (Fig. 4 A and B and Fig. S7). Collectively, these observations suggest that, contrary to what was previously thought, all ZP modules share a common architecture, so that other molecular features must regulate polymerization specificity.
Fig. 3.
Fig. 4.
Fig. S6.
Fig. S7.
A Structured Interdomain Linker Is Conserved in Self-Polymerizing ZP Modules.
Structure comparison reveals a striking difference in the linker between ZP-N and ZP-C domains: Whereas this region is highly flexible in ZP3 (3), UMOD contains a rigid linker formed by α1 and β1 before the IHP (Fig. 4B and Fig. S5B). Analysis of UMOD ZP-C truncation constructs indicates that both of these secondary structure elements, which are also present in GP2, TECTA, and ZPD (Fig. S3), are essential for folding and secretion (Fig. 4C). Whereas UMOD430–610 starting with α1 is secreted comparably to UMODp (Fig. 4C, lanes 1–2), constructs beginning with β1 (UMOD440–610) or βA (UMOD451–610) are almost completely retained in the cell (Fig. 4C, lanes 3–4 and 9–10).
Like ZP3, ZP2 contains a ZP-N/ZP-C linker (Fig. S8); however, although this region was present in the crystals (Fig. S6C), ZP2 ZP-C is only defined from the IHP onward (Fig. 4B and Figs. S6D and S7). Moreover, unlike in the case of UMOD, the linker is not required for secretion of ZP2 ZP-C (Fig. 4C, lanes 5–6).
Fig. S8.
UMOD linker α1 packs tightly against the IHP-containing β-sheet (Fig. 4D and Fig. S7), shielding from the solvent hydrophobic residues also found in GP2, TECTA, and, to a lesser extent, ZPD (Fig. S3). Mutation of conserved α1 residues D430 and L435 causes trafficking and assembly defects of UMOD (10), whereas changes affecting amino acids located on the opposite side (A461E and G488R) are associated with kidney disease (Fig. S3). Thus, UMOD function is compromised upon disruption of contacts between α1/β1 and ZP-C. This interaction constrains the relative orientation between ZP-N and ZP-C, so that UMOD adopts an extended conformation that is significantly different from the conformation of ZP3 (Fig. 5). In the latter, as well as in ZP2, the linker lacks α1/β1 and the IHP-containing β-sheet surface is hydrophilic, resulting in a compact arrangement wherein ZP-N folds back onto ZP-C.
Fig. 5.
ZP-N Domain Dimerization Is Required for UMOD Polymerization.
A major consequence of the extended configuration of the ZP module of UMOD is that the hydrophobic surface formed by ZP-N βA/βG is free to dimerize with the same region of a neighboring ZP-N through parallel β-sheet extension, burying a surface area of 2,148 Å2 (Figs. 2A and 6A). Computational analysis using PISA (34) scores this ZP-N/ZP-N interface as highly significant, and inward-facing hydrophobic residues in βA/βG are conserved across UMOD, GP2, TECTA, and ZPD (Fig. S3). Furthermore, the interface involves the N396 glycan, which forms intermolecular hydrogen bonds with the other UMOD molecule (Fig. 6A) and is also conserved among filament-forming ZP modules (Fig. S3). Notably, mutation of the corresponding N-glycosylation site of TECTA is associated with hearing loss (35), suggesting that this carbohydrate is important for tectorial membrane assembly.
Fig. 6.
To evaluate the biological significance of the ZP-N homodimer, conserved interface residues (Fig. S3) were individually mutated to Lys to prevent edge-to-edge β-sheet interaction (36). Whereas mutation of peripheral residues L329 and I419 does not significantly affect UMOD assembly (Fig. S9 A–C), mutation of core residues L333 and I421 (Fig. 6A) completely abolishes filament formation (Fig. 6 D and E) compared with WT UMOD (Fig. 6 B and C). Accordingly, EM of corresponding mMBP-fused mutants of L333 and I421 detects no filaments (Fig. S9 D–F). Considering that neither mutation affects the trafficking (Fig. S9 G–I), secretion (Fig. S9J), or proteolysis (Fig. S9K) of UMOD, we conclude that the homodimer observed in our crystals represents a polymerization intermediate, whose formation is essential for the assembly of UMOD filaments.
Fig. S9.
Sequence alignments and structural data indicate that the two moieties of the ZP module can be joined by very few residues (BG and ENG) or connected by a linker that is either unstructured (ZP1–ZP4, and TECTB) or structured (UMOD, GP2, and TECTA) (Figs. S3 and S8). Remarkably, these combinations coincide with the different polymerization abilities of the corresponding proteins: BG and ENG do not polymerize; ZP1–ZP4 and TECTB heteropolymerize; and UMOD, GP2, and TECTA homopolymerize (7, 17, 37, 38). This observation is consistent with the idea that in the last set of proteins, coupling of an α1/β1-containing linker to ZP-C induces an extended conformation of the ZP module. This conformation, in turn, exposes the βA/βG surface of ZP-N to form a dimer that initiates homopolymerization. On the other hand, the presence of a flexible linker may allow ZP1–ZP4 to adopt a secretion-competent conformation, such as the conformation observed in the structure of full-length ZP3 (3), which could require additional factors to trigger heteropolymerization and incorporation into the egg coat (39).
Conclusion
First isolated more than 60 years ago (40) and redescribed 35 years later as UMOD (41), UMOD has been recognized as a guardian against urinary tract infection and a crucial player in innate immunity; kidney disease; and, more recently, hypertension (1, 2, 42, 43). Our work gives mechanistic insights into how UMOD and other ZP module proteins assemble into their biologically active form, and how their structure and polymerization can be perturbed by pathogenic human mutations.
Materials and Methods
For structural studies, mMBP-UMODpXR and ZP2 ZP-C proteins were transiently expressed in HEK293S and HEK293T cells, respectively, based on published protocols (44–46); immunofluorescence studies were performed using stably transfected MDCK cell lines, essentially as described (10). Construct information and detailed methods for protein purification, deglycosylation, crystallization, and structure determination; UMOD filament preparation; and EM and immunofluorescence analyses are provided in SI Materials and Methods. X-ray data collection and refinement statistics are summarized in Table S1. Atomic coordinates and structure factors for human UMODpXR and mouse ZP2 ZP-C have been deposited in the Protein Data Bank (ID codes 4WRN and 5BUP, respectively). Urine for EM analysis of native UMOD was kindly donated by M. Bokhove.
SI Materials and Methods
DNA Constructs.
Expression constructs were generated by PCR using PfuTurbo DNA polymerase (Agilent Technologies); mutations were introduced by overlap extension PCR or with a QuikChange Site-Directed Mutagenesis Kit (Agilent Technologies). Oligonucleotides were from Sigma–Aldrich, and all constructs were verified by DNA sequencing (Eurofins Genomics).
Except for immunofluorescence studies, which relied on previously described plasmids encoding HA-tagged WT UMOD and CCS4A mutants (586-RFRS-589 to AAAA) (10), all constructs were based on mammalian expression vector pHLsec (44). For expression of secreted fusions, a codon-optimized gene encoding an N-terminally 6His-tagged mMBP was synthesized (GenScript), which includes a combination of mutations increasing maltose affinity and crystallizability (47–50). The mMBP gene was cloned in-frame to the chicken CRYPα1 R2G signal peptide-encoding sequence of pHLsec, and includes a 3′ NotI restriction site that results in a three-Ala linker between mMBP and passenger proteins. Nonpolymerizing constructs for X-ray crystallographic studies were generated using cDNA fragments carrying mutations of the CCS-encoding sequences of human UMOD (R586A/R588A) and mouse ZP2 (K634N/R635A). UMOD N-glycosylation site N513, which is not essential for secretion, was also mutated in UMODpXR, as well as the UMOD truncation mutants analyzed in Fig. 4C.
Protein Expression.
DNA for small- and large-scale transfections was prepared using HiSpeed Plasmid Midi Kits and EndoFree Plasmid Maxi/Giga Kits, respectively (both from QIAGEN). HEK293 cells were cultivated using DMEM supplemented with 4 mM l-glutamine (Life Technologies) and 10% (vol/vol) FBS (Saveen & Werner) at 37 °C in 5% CO2. Subsequently, they were transiently transfected in DMEM with 4 mM l-glutamine using 25-kDa branched polyethylenimine (Sigma–Aldrich), essentially as described (44). Small-scale transfections were performed in six-well plates (10 cm2 per well; Corning), large-scale transfections were carried out in T-flasks (150 cm2; BD Biosciences) or ribbed roller bottles (2,125 cm2; Greiner).
Production of glycosylated mMBP-UMODpXR was performed using HEK293S cells (CRL-3022; American Type Culture Collection), which secrete homogeneous Man5GlcNAc2-glycosylated proteins that can be treated with endoglycosidase H (Endo H) to remove N-glycans (45, 46). ZP2 ZP-C, which does not contain N-linked glycosylation sites, was expressed in HEK293T cells. In both cases, conditioned medium was harvested 3 d after transfection. Notably, fusion to mMBP was absolutely essential to obtain suitable amounts of well-behaved UMODp, as well as for its crystallization.
MDCK cell lines stably expressing HA-tagged constructs of UMOD were generated as described previously, and conditioned media and cell lysates were prepared as published (10).
mMBP-UMODp filaments were produced by transiently cotransfecting HEK293T cells with constructs expressing WT or mutant mMBP-fused UMODS292–Q640 in a 10:1 molar ratio with a pcDNA3.1(+)-derived vector expressing human hepsin (8).
Protein Analysis.
Cell lysates were obtained by resuspending cells from a 10-cm2 culture well in lysis buffer [50 mM Tris⋅HCl/50 mM 3-(N-morpholino)propanesulfonic acid (pH 7.7), 0.1% SDS, 1 mM EDTA] supplied with protease inhibitors (Roche), followed by centrifugation for 10 min at 18,000 × g at 4 °C and filtration using a 0.22-μm syringe filter (Millipore).
Samples were separated on SDS/PAGE gels and transferred to nitrocellulose membranes (GE Healthcare). Immunoblotting was performed with Penta-His mouse mAb (1:1,000; QIAGEN) or anti-HA mouse Ab (1:1,000; Covance). Chemiluminescence detection was performed with Western Lightning ECL Plus (PerkinElmer) or using an Immobilon Western Chemiluminescent Horseradish Peroxidase Substrate Kit (Millipore).
Protein Purification and Deglycosylation.
Conditioned medium was adjusted to 5 mM imidazole, 150 mM NaCl, 20 mM Na-Hepes (pH 8.0) [immobilized metal affinity chromatography (IMAC) binding buffer]. Ten mL of preequilibrated nickel-nitrilotriacetic acid (Ni-NTA) agarose slurry (QIAGEN) was then added per L of medium and allowed to incubate overnight at 4 °C on a shaker. Ni-NTA beads were collected, washed with IMAC binding buffer, and batch-eluted with 500 mM imidazole, 150 mM NaCl, and 20 mM Na-Hepes (pH 8.0). The IMAC elution fraction was concentrated using centrifugal filtration devices (Amicon) with an appropriate molecular weight cutoff (MWCO). In the case of mMBP-UMODpXR expressed in HEK293S cells, concentrated fusion protein was deglycosylated with Endo H (1:10 mass ratio) for 1 h at 37 °C in 120 mM Na/K phosphate (pH 6.0). Concentrated material was applied to a Superdex 200 26/600 size exclusion chromatography (SEC) column attached to an ÄKTAFPLC system (GE Healthcare) and preequilibrated with 100 mM NaCl, 20 mM Na-Hepes (pH 8.0), and 10 mM d-maltose. For purification of ZP2 ZP-C, all buffers contained 500 mM NaCl and a Superdex 75 26/600 column was used. SEC fractions containing purified proteins were pooled, concentrated, and used for crystallization trials.
Protein Crystallization.
Purified mMBP-UMODpXR was concentrated to 6.5–15.0 mg/mL in 100 mM NaCl, 20 mM Na-Hepes (pH 8.0), and 1.5 mM maltose and was crystallized at room temperature (RT) by hanging drop vapor diffusion against mother liquor containing 900 mM Na/K tartrate and Tris⋅HCl (pH 7.0–8.6) (Fig. S1A). Crystals could only be obtained in the presence of Zn(OAc)2, which was added in a 1:1.5 molar ratio to the mMBP-UMODpXR solution before crystallization and was eventually found to mediate crystal packing by interacting with the 6His-tag of mMBP-UMODpXR. Crystals grew to 200–400 μm and were subsequently slowly accommodated to 4 °C. Finally, they were transferred in six steps to mother liquor containing 20–30% (vol/vol) glycerol and flash-cooled in liquid nitrogen before data collection at 100 K.
Purified ZP2 ZP-C was concentrated to 5.0 mg/mL in 20 mM Na-Hepes (pH 8.0) and 200 mM NaCl and was crystallized at RT by hanging drop vapor diffusion against 25% (wt/vol) PEG 3350, 100 mM NaOAc (pH 5.5), 200 mM (NH4)2SO4, and 20% (vol/vol) glycerol. Microseeding was necessary to grow single crystals of adequate size for analysis, and produced well-ordered crystals up to 0.5 mm long (Fig. S6A). Crystals were flash-cooled in liquid nitrogen before data collection at 100 K.
X-Ray Diffraction Data Collection.
Native datasets were collected using PILATUS 6M-F detectors at European Synchrotron Radiation Facility beamline ID29 (51) (mMBP-UMODpXR) or Diamond Light Source beamline I02 (ZP2 ZP-C). Data collection statistics can be found in Table S1. Anomalous difference data were obtained from multiple passes collected at a wavelength of 1.8 Å.
Data Processing and Structure Determination.
All datasets were integrated and scaled with XDS (52).
The structure of mMBP-UMODpXR was solved by molecular replacement (MR) with PHASER (53), using MBP coordinates extracted from PDB ID code 3D4G (5) as a search model. Successful MR runs showed a clear positive different density peak in the MBP binding site for maltose, which was not included in the search model. After obtaining initial MR phases, density modification was performed with RESOLVE (54) in the PHENIX package (55). The model of mMBP-UMODpXR was generated by combining multiple rounds of PHENIX AutoBuild (56) with Buccaneer (57) and manual building in Coot (58).
Structure Analysis.
Structure comparisons were performed using PDBeFold (63). Oligomeric state was analyzed using PISA (34). PISA analysis shows that ZP-N/ZP-N dimer formation buries 16.3% and 9.0% of the total ZP-N domain and ZP module surface, respectively, resulting in a ΔG of −11.6 kcal/mol. Together with a complexation significance score of 1.0 and the lack of any additional interaction scored as significant, this analysis suggests that the interface observed in the crystals is biologically relevant. Furthermore, the relative orientation of mMBP and UMODpXR is different in the two molecules, indicating that the mMBP fusion is flexible and mMBP does not affect the overall structure of UMODpXR itself.
UMOD Filament Preparation.
Native UMOD filaments were purified from healthy male urine as described (64), and elastase-resistant filaments were prepared as previously reported (18).
To purify recombinant filaments, 1 mL of conditioned medium of UMOD-expressing HEK293T cells was harvested and spun for 5 min at 500 × g to remove cell debris. The supernatant was dialyzed at 4 °C against 10 mM Na-Hepes (pH 8.0) and 10 mM NaCl using a 30-kDa MWCO Slide-A-Lyzer dialysis cassette (Thermo Scientific). After dialysis, the material was centrifuged for 30 min at 18,000 × g to remove aggregates, followed by the addition of 150 mM NaCl to the supernatant and incubation on ice for 1 h to gelify UMOD filaments. The supernatant was then spun for 2 h at 18,000 × g, resulting in a small pellet of filaments that was finally vigorously resuspended in 20 μL of ultrapure water.
EM.
The sample (5 μL) was applied to a glow-discharged 400 mesh copper grid with a carbon support film. After 2 min, the solution was removed with filter paper, followed by washing with 20 mM Tris⋅HCl (pH 7.5) and 150 mM NaCl. A final wash step was performed with ultrapure water, followed by negative staining with 2% (wt/vol) uranyl acetate. Samples were analyzed using a CM120 electron microscope (Philips) equipped with a LaB6 electron source. Images were recorded on a 1K Tietz camera or with SO-163 electron film (Kodak). Films were digitized using an Epson Perfection 4990 PHOTO flatbed scanner.
For on-grid immunogold labeling, 5 μL of diluted filament solution was applied to glow-discharged 400 mesh gold grids with a carbon support film. After 5 min, the grids were washed by inverting them onto a drop of ultrapure water for 2 min and then moved onto a 30-μL drop of blocking buffer [20 mM Tris⋅HCl (pH 7.5), 150 mM NaCl, 0.1% BSA, or 0.25% blotting grade nonfat dry milk] for 15 min. Grids were then moved onto a 10-μL drop of blocking buffer containing a 1:20 or 1:40 dilution of anti-HA Ab [HA.11 clone 16B2 purified mouse monoclonal primary Ab (Covance)] for 1 h. Grids were washed by inverting them three times for 5 min onto 30-μL drops of washing buffer [20 mM Tris⋅HCl (pH 7.5), 150 mM NaCl] and moved onto a 20-μL drop of blocking buffer containing a 1:40 dilution of goat anti-mouse Ab [18 nm of gold-conjugated purified goat polyclonal Ab against mouse IgG-Fc (Abcam)] for 1 h. After washing as previously detailed, followed by three 5-min washes in ultrapure water, grids were negatively stained and imaged as described above.
Immunofluorescence Analysis.
Immunofluorescence studies were performed essentially as described (10). MDCK cells grown on coverslips were fixed in 4% (wt/vol) paraformaldehyde for 20 min at RT. When needed, cells were permeabilized for 20 min at RT with PBS solution-0.5% (vol/vol) Triton X-100 [note that this procedure essentially removes UMOD filaments from the cell surface (65)]. Permeabilized or unpermeabilized cells were incubated with 10% (vol/vol) preimmune donkey serum (Abcam) for 30 min at RT and then labeled for 1 h at RT with anti-UMOD primary Ab diluted in PBS with 1% (vol/vol) donkey serum (1:500; MP Biomedicals). Cells were washed in PBS and incubated with the appropriate Alexa Fluor 594-conjugated secondary Ab [1:500 in PBS with 1% (vol/vol) donkey serum; Life Technologies]. Nuclei were stained with DAPI (Life Technologies), and slides were mounted using fluorescent mounting medium (DAKO). Slides were visualized with a DM 5000B fluorescence upright microscope (Leica DFC480 camera, Leica DFC Twain Software, 40×/0.75 lens; Leica Microsystems). All images were imported in Photoshop CS (Adobe Systems) and adjusted for brightness and contrast.
Sequence Analysis.
Except for D8C (66), EGF IV, and ZP-N/ZP-C (this study), domain boundaries reported in Fig. 1A were based on SMART (67). The sequence alignment reported in Fig. S3 was generated using T-Coffee (68) and assembled using ESPript (69). The alignment in Fig. S8 was generated with PROMALS3D (70), manually edited, and colored using TEXshade (71). Cys covariation analysis was performed by manual inspection of ZP-like domain (PF00100) seed sequence alignments from Pfam (32).
Data Availability
Data deposition: Atomic coordinates and structure factors have been deposited in the Protein Data Bank, www.pdb.org (PDB ID codes 4WRN and 5BUP).
Acknowledgments
This work is dedicated to the memory of Franca Serafini-Cessi, whose studies increased our understanding of uromodulin biology. Correspondence with Dr. Serafini-Cessi directly inspired this work. We thank M. Monné (Università degli Studi della Basilicata) for initial work on the project, H. Hebert and the Department of Biosciences and Nutrition (Karolinska Institutet) for access to the Center for High Resolution Electron Microscopy, the European Synchrotron Radiation Facility (ESRF; Grenoble) and Diamond Light Source (DLS; Oxford) for beam time (ESRF: mx1416/mx1551/mx1639, DLS: mx8492-18/mx8492-34), and G. Wallis (University of Otago) for comments and discussion. We are grateful to R. Aricescu and Y. Zhao (University of Oxford) for mammalian expression vector pHLsec and HEK293T cells, to D. Leahy (Johns Hopkins University School of Medicine) for Escherichia coli expression vector pProEX HT-endoglycosidase H, and to D. Waugh (National Cancer Institute) for E. coli strain BL21(DE3)-RIL/pRK793. This research was supported by the Karolinska Institutet, the Center for Innovative Medicine, Swedish Research Council Grant 2012-5093, the Göran Gustafsson Foundation for Research in Natural Sciences and Medicine, the Sven and Ebba-Christina Hagberg Foundation, a European Molecular Biology Organization Young Investigator award, the European Research Council (ERC) under the European Union’s Seventh Framework Programme (FP7/2007-2013)/ERC Grant Agreement 260759 (to L.J.); and the Fondazione Telethon (GGP14263), the Italian Ministry of Health (RF-2010-2319394), and Fondazione Cariplo (2014-0827) (to L.R.). Crystallographic data collection was also supported by FP7/2007-2013 under BioStruct-X (Grant Agreement 283570).
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Information & Authors
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Freely available online through the PNAS open access option.
Data Availability
Data deposition: Atomic coordinates and structure factors have been deposited in the Protein Data Bank, www.pdb.org (PDB ID codes 4WRN and 5BUP).
Submission history
Published online: January 25, 2016
Published in issue: February 9, 2016
Keywords
Acknowledgments
This work is dedicated to the memory of Franca Serafini-Cessi, whose studies increased our understanding of uromodulin biology. Correspondence with Dr. Serafini-Cessi directly inspired this work. We thank M. Monné (Università degli Studi della Basilicata) for initial work on the project, H. Hebert and the Department of Biosciences and Nutrition (Karolinska Institutet) for access to the Center for High Resolution Electron Microscopy, the European Synchrotron Radiation Facility (ESRF; Grenoble) and Diamond Light Source (DLS; Oxford) for beam time (ESRF: mx1416/mx1551/mx1639, DLS: mx8492-18/mx8492-34), and G. Wallis (University of Otago) for comments and discussion. We are grateful to R. Aricescu and Y. Zhao (University of Oxford) for mammalian expression vector pHLsec and HEK293T cells, to D. Leahy (Johns Hopkins University School of Medicine) for Escherichia coli expression vector pProEX HT-endoglycosidase H, and to D. Waugh (National Cancer Institute) for E. coli strain BL21(DE3)-RIL/pRK793. This research was supported by the Karolinska Institutet, the Center for Innovative Medicine, Swedish Research Council Grant 2012-5093, the Göran Gustafsson Foundation for Research in Natural Sciences and Medicine, the Sven and Ebba-Christina Hagberg Foundation, a European Molecular Biology Organization Young Investigator award, the European Research Council (ERC) under the European Union’s Seventh Framework Programme (FP7/2007-2013)/ERC Grant Agreement 260759 (to L.J.); and the Fondazione Telethon (GGP14263), the Italian Ministry of Health (RF-2010-2319394), and Fondazione Cariplo (2014-0827) (to L.R.). Crystallographic data collection was also supported by FP7/2007-2013 under BioStruct-X (Grant Agreement 283570).
Notes
This article is a PNAS Direct Submission. P.W. is a guest editor invited by the Editorial Board.
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The authors declare no conflict of interest.
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