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RESEARCH ARTICLE

Different roles of cadherins in the assembly and structural integrityof the desmosome complex

Molly Lowndes1, Sabyasachi Rakshit2,3, Omer Shafraz2,3, Nicolas Borghi4, Robert M. Harmon5,Kathleen J. Green5, Sanjeevi Sivasankar2,3 and W. James Nelson1,6,7,*

ABSTRACT

Adhesion between cells is established by the formation of specialized

intercellular junctional complexes, such as desmosomes.

Desmosomes contain isoforms of two members of the cadherin

superfamily of cell adhesion proteins, desmocollins (Dsc) and

desmogleins (Dsg), but their combinatorial roles in desmosome

assembly are not understood. To uncouple desmosome assembly

from other cell–cell adhesion complexes, we used micro-patterned

substrates of Dsc2aFc and/or Dsg2Fc and collagen IV; we show that

Dsc2aFc, but not Dsg2Fc, was necessary and sufficient to recruit

desmosome-specific desmoplakin into desmosome puncta and

produce strong adhesive binding. Single-molecule force

spectroscopy showed that monomeric Dsc2a, but not Dsg2, formed

Ca2+-dependent homophilic bonds, and that Dsg2 formed Ca2+-

independent heterophilic bonds with Dsc2a. A W2A mutation in

Dsc2a inhibited Ca2+-dependent homophilic binding, similar to

classical cadherins, and Dsc2aW2A, but not Dsg2W2A, was

excluded from desmosomes in MDCK cells. These results indicate

that Dsc2a, but not Dsg2, is required for desmosome assembly

through homophilic Ca2+- and W2-dependent binding, and that Dsg2

might be involved later in regulating a switch to Ca2+-independent

adhesion in mature desmosomes.

KEY WORDS: Desmosome, Adhesion, Desmocollin, Desmoglein

INTRODUCTIONCell–cell adhesion is essential for the organization andmaintenance of complex tissues in multicellular organisms. Thecadherin superfamily of cell–cell adhesion proteins is essential

for tissue formation throughout the Metazoa (Franke, 2009; Odaand Takeichi, 2011). Disruption of cadherin binding generallyresults in tissue defects, and is common in metastatic cancers(Chidgey and Dawson, 2007; Garrod, 1995; Runswick et al.,

2001; Saito et al., 2012; Simpson et al., 2011). There are fourmajor subfamilies of cadherins: the classical cadherins,desmosomal cadherins, protocadherins and atypical cadherins

(Shapiro and Weis, 2009).

Desmosomes comprise two different cadherins, termed

desmogleins (Dsg) and desmocollins (Dsc), that form a protein-

dense adhesion complex that is required in tissues and organs that

resist mechanical stress (Green and Simpson, 2007; Green et al.,

1998). For example, the intercalated discs that connect heart

muscle cells contain one isoform of each subtype (Dsc2 and

Dsg2) (Delva et al., 2009), and mutations in Dsc2 and Dsg2 cause

hereditary diseases such as arrhythmogenic right ventricular

cardiomyopathy (ARVC) (Bhuiyan et al., 2009; Heuser et al.,

2006) and mild striate palmoplantar keratoderma (SPPK)

(Simpson et al., 2009). Several of these hereditary mutations

have been mapped to the extracellular domain of Dsg2 and Dsc2

(Gaertner et al., 2012; Gehmlich et al., 2011). In the epidermis,

different Dsg and Dsc isoforms are expressed in a cell-type-

specific manner, and autoimmune diseases such as pemphigus

foliaceus and pemphigus vulgaris target extracellular binding of

Dsg1 and Dsg3, causing loss of adhesion in the epidermis and

mucosal membranes (Amagai and Stanley, 2012; Waschke,

2008). The range of heart and skin defects caused by disruption

of desmosomal adhesion suggests that Dsc and Dsg play

important roles in cell–cell adhesion and differentiation (Kottke

et al., 2006).

Desmosomal cadherins contain evolutionarily conserved

domains, with low sequence similarity (,30%) to classical

cadherins, that include five extracellular ‘cadherin’ repeats, a

transmembrane domain and a cytoplasmic domain that binds

desmosome-specific adaptor proteins (Patel et al., 2003).

Desmosomes form punctate structures at cell–cell contacts

(Pasdar and Nelson, 1988a; Pasdar and Nelson, 1988b) that

comprise an intercellular midline corresponding to sites of trans

interactions between opposing cadherins, and a dense plaque

structure in the cytoplasm that connects to intermediate filaments

(Al-Amoudi et al., 2011; Al-Amoudi et al., 2007; He et al., 2003;

North et al., 1999).

Analysis of the structure of classical cadherins bound in trans

identified a crucial tryptophan at position 2 (W2) in the first N-

terminal extracellular (EC1) repeat that forms a ‘strand swap

dimer’ with the opposing cadherin (Boggon et al., 2002). An

alanine replacement mutation (W2A), which inhibits strand swap

dimerization, revealed a second configuration termed the X-

dimer, which involves residues near the EC1–EC2 Ca2+-binding

sites, and is thought to be an intermediate that facilitates the

formation of the W2 strand-swap dimer (Harrison et al., 2010;

Sivasankar et al., 2009). Importantly, homophilic binding of Dsc2

is blocked by mutations in W2 and A80, the latter of which

contributes to the hydrophobic pocket into which W2 inserts

during strand swapping (Nie et al., 2011), but there is no sequence

similarity in desmosomal cadherins to residues required for the

X-dimer.

1Cancer Biology Program, Stanford University, Stanford, CA 94305, USA.2Department of Physics and Astronomy, Iowa State University, Ames, IA 50011,USA. 3Ames Laboratory, United States Department of Energy, Ames, IA 50011,USA. 4Institut Jacques Monod, Unite Mixte de Recherche 7592, Centre Nationalde la Recherche Scientifique, and Universite Paris-Diderot, 75013 Paris, France.5Department of Pathology, Northwestern University Feinberg School of Medicine,Chicago, IL 60611, USA. 6Department of Biology, Stanford University, Stanford,CA 94305, USA. 7Department of Molecular and Cellular Physiology, StanfordUniversity, Stanford, CA 94305, USA.

*Author for correspondence (wjnelson@stanford.edu)

Received 15 November 2013; Accepted 4 February 2014

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The cytoplasmic domains of Dsc isoforms are derived fromtwo different splicing events (a, b), whereas the cytoplasmic

domains of Dsg isoforms are longer and might have additionalfunctions (Brennan and Mahoney, 2009; Brennan et al., 2007;Getsios et al., 2009; Harmon et al., 2013). Biochemical studiesindicate that the cytoplasmic domain of both Dsc and Dsg have

binding sites for the core desmosomal adaptor proteinplakoglobin (Kowalczyk et al., 1996; Troyanovsky et al.,1994b), which binds other desmosome-specific proteins,

plakophilin (PKP) and desmoplakin (DP I/II, the two majorproteins encoded by DSP), that link the complex to intermediatefilaments (cytokeratins) (Bornslaeger et al., 1996; Kowalczyk

et al., 1997; Troyanovsky et al., 1994a). There is also evidencethat DP I/II can be recruited to Dsc–Dsg complexes in theabsence of plakoglobin (Troyanovsky et al., 1994a; Troyanovsky

et al., 1994b). The recruitment of these cytoplasmic adaptorproteins and the intermediate filament cytoskeleton is thought topromote Dsc–Dsg clustering and stabilize the structural integrityof the desmosome.

Both Dsg and Dsc are thought to be involved in extracellulartrans interactions between adjacent cells (Chitaev andTroyanovsky, 1997; Dusek et al., 2007; Getsios et al., 2004;

Marcozzi et al., 1998; Roberts et al., 1998; Runswick et al.,2001). How Dsc and Dsg combine to form adhesive bonds isunclear because different binding properties have been assigned

to the most ubiquitously expressed isoforms (Dsg2 and Dsc2)depending on the assay used (Waschke, 2008). Knockdown ofDsc2 (Roberts et al., 1998) or knockout of Dsg2 expression

(Eshkind et al., 2002) results in a loss of functional desmosomes.In addition, Dsg2 and Dsc2 can be depleted from junctions byblocking their microtubule (MT)-mediated trafficking, eachresulting in weakened adhesive strength (Nekrasova et al.,

2011). Cell-free studies showed that the EC1–EC2 domain ofDsc2 can form homophilic bonds and heterophilic bonds withDsg2, but the EC1–EC2 domain of Dsg2 does not form

homophilic bonds (Syed et al., 2002). In contrast, atomic forcemicroscope (AFM) experiments with dimeric Fc fusion proteinshave indicated that Dsg2 can also form homophilic bonds

(Hartlieb et al., 2013; Schlegel et al., 2010). Cell-based assays inkeratinocytes using cross-linking reagents found only homophilicbinding for both Dsg2 and Dsc2 (Nie et al., 2011). Thus, therequirement for both Dsg and Dsc in the structural organization

and maintenance of the desmosomal complex remains unclear.To distinguish the roles of Dsc2a and Dsg2, we used three

independent assays: a desmosome assembly assay using dual-

patterned surfaces containing purified Dsg2, Dsc2a, or acombination of Dsc2a plus Dsg2, and collagen IV; a single-molecule binding assay with wild-type and W2A mutant

monomeric Dsg2 and Dsc2a using single-molecule forcespectroscopy (SMFS); and a cell-based assay to examine theorganization and stability of wild-type and W2A mutant Dsg2

and Dsc2a in desmosomes in situ. Our results identify uniqueroles of Dsc2a and Dsg2 in desmosome organization.

RESULTSDifferent requirements of desmosomal cadherins for theformation of punctate desmosomal structuresTo distinguish roles of Dsg2 and Dsc2a in desmosome adhesion

and assembly, we adapted a method (Borghi et al., 2010) toanalyze cell binding to micro-patterned surfaces of Dsc2a and/orDsg2, and the extracellular matrix protein collagen IV. This assay

uncouples desmosome assembly from other cell adhesion

complexes such as E-cadherin. The complete extracellulardomain (EC1–5) of Dsg2 and Dsc2a was fused to the constant

region of human IgG1 (Fc) (supplementary material Fig. S1A), andDsg2Fc and Dsc2aFc were expressed in mammalian HEK293Tcells to ensure correct post-translational modifications. Fusionproteins were secreted into the medium, purified by affinity

chromatography with Protein-A–Sepharose (Drees et al., 2005),and labeled with Cy5 dye to visualize protein patterns on micro-patterned coverslips (supplementary material Fig. S1B).

Patterned 10 mm stripes were contact printed with collagen IV,followed by saturating amounts of Dsg2Fc, Dsc2aFc or Dsg2Fcplus Dsc2aFc (50:50) so that the collagen IV stripes were

juxtaposed to Dsc2a/Dsg2 stripes, with both stripes showinguniform staining of the proteins (supplementary material Fig.S1C). MDCK cells were seeded onto the dual-patterned

coverslips and allowed to adhere for at least 2 h beforefixation. Total internal reflection fluorescence microscopy(TIRF-M) was used to detect cellular DP I/II on the ventralmembrane of MDCK cells attached to stripes containing Dsg2Fc,

Dsc2aFc, or Dsg2Fc plus Dsc2aFc (Fig. 1). Given that MDCKcells express both Dsc2a and Dsg2 (Pasdar and Nelson, 1988a;Pasdar and Nelson, 1988b) they could form homophilic and/or

heterophilic bonds with Dsc2aFc and Dsg2Fc stripes and recruitother desmosomal complex proteins such as DP I/II, which linksthe complex to the cytoskeleton.

Cellular DP I/II was distributed evenly over alternating stripesof collagen IV and Fc (Fig. 1A). In contrast, cellular DP I/IIdistribution was restricted over stripes containing Dsg2Fc

(Fig. 1B), Dsc2aFc (Fig. 1C) or Dsg2Fc plus Dsc2aFc(Fig. 1D), and generally excluded from areas over the collagenIV stripe. Significantly, the organization of DP I/II was diffuseover the Dsg2Fc stripe (Fig. 1B), but more punctate over the

Dsc2aFc stripe (Fig. 1C) and Dsg2Fc plus Dsc2aFc stripe(Fig. 1D). Indeed, the DP I/II puncta over Dsg2Fc plusDsc2aFc stripes appeared similar to bone fide desmosomes

between adherent pairs of MDCK cells (supplementary materialFig. S1D). The punctate localization of DP I/II does not appear tobe due to a punctate distribution of Dsc2aFc or Dsg2Fc in the

stripes, as both of them were evenly distributed (supplementarymaterial Fig. S1E). These results indicate that Dsc2a is necessaryfor DP I/II organization into desmosome-like puncta, whereasDsg2 is not required on at least one of the opposing surfaces.

Desmosomal cadherins have distinct binding propertiesThe results from MDCK binding to micro-patterned surfaces

indicate that formation of punctate desmosome-like cellularstructures involved homophilic binding of Dsc2a, but not Dsg2.Heterophilic interactions, if they occurred, were not sufficient to

form these puncta. To directly address whether Dsc2a and Dsg2form homophilic and/or heterophilic interactions, we measuredtheir binding using SMFS with an AFM. We did not use the Fc

fusion construct in the SMFS experiments because Fcdimerization positions the cadherins in close proximity in a cisorientation (Zhang et al., 2009); thus, interactions betweenopposing cadherin pairs cannot be distinguished from single-

molecule interactions which might introduce artifacts in datainterpretation. Therefore, we designed monomeric Dsc2a andDsg2 fusion proteins, as described previously for E-cadherin

(Rakshit et al., 2012; Sivasankar et al., 2009; Zhang et al., 2009),comprising the extracellular domain of Dsc2a or Dsg2 fused atthe C-terminal to a tag comprising an Avi tag, for binding biotin

attached to a surface, and a TEV-removable His tag, for

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purification from HEK293T cell conditioned medium using a Ni-NTA column (pATH; Fig. 2A). The purified proteins had anexpected molecular mass of ,90 kDa, similar to the E-cadherin

extracellular domain fused to the same tag (Fig. 2B); a slowermigrating protein was detected occasionally and is likely theuncleaved precursor (Fig. 2B, *).

Purified Dsg2pATH and Dsc2apATH monomers were

biotinylated and immobilized via polyethylene glycol (PEG)linkers to a glass coverslip and an AFM cantilever tip (Fig. 2C),as described previously for E-cadherin (Rakshit et al., 2012;

Sivasankar et al., 2009; Zhang et al., 2009). The AFM tip andcoverslip were first brought into contact so that opposing cadherinsinteracted. The tip was then withdrawn from the surface so that

force was applied to the bond. Single-molecule interactions wereidentified from the freely-jointed chain stretching of the polymertether that anchored the proteins to the surface (Smith et al., 1992)(Fig. 2D); the contour length of the stretched polymer was used to

distinguish specific interactions from non-specific binding (Zhanget al., 2009). Approximately 1000–2000 measurements werecarried out at 11 different pulling rates (loading rates), and

histograms of the unbinding forces were compiled (supplementarymaterial Fig. S2). Dsg2pATH and Dsc2apATH densities on theAFM tip and coverslip were adjusted to yield binding frequencies

in the 5% range (Poisson statistics predicts that more than 95% ofthe measured events at this unbinding frequency occur due tosingle-molecule unbinding), and the adjusted densities were kept

constant for all experimental conditions.The force required to rupture the bond between homophilic

(Dsg2pATH–Dsg2pATH or Dsc2apATH–Dsc2apATH) or heterophilic(Dsg2pATH–Dsc2apATH) interactions was measured in the presence

of Ca2+ or EGTA, to chelate Ca2+ (Fig. 2E). Non-specific binding

rates were measured using identically processed AFM cantilevers andcoverslips that were not decorated with Dsg2pATH or Dsc2apATH.The binding frequency (percentage) showed that the greatest number

of unbinding events involved homophilic interactions betweenDsc2apATH (.4%), and that binding was significantly reduced, to,2%, upon addition of EGTA, indicating a Ca2+-dependent bindingmechanism. In contrast, the homophilic binding frequency between

Dsg2pATH was ,1%, which was below that of non-specific bindingevents (Fig. 2E). We detected heterophilic binding events betweenDsc2apATH and Dsg2pATH, and the frequency of those binding

events was similar in the presence or absence of Ca2+, indicating aCa2+-independent binding mechanism (Fig. 2E). Given thatDsg2pATH bound to Dsc2apATH through heterophilic interactions,

the lack of homophilic Dsg2pATH binding is not due to inactivity ofDsg2pATH.

Because the unbinding force increases linearly with the naturallog of the loading rate (Fig. 2F), we used the Bell–Evans model

(Eqn 3) (Bell, 1978; Evans and Ritchie, 1997) to determineintrinsic unbinding rates. Our results showed that the unbindingrates of Ca2+-dependent homophilic binding of Dsc2apATH and

Ca2+-independent heterophilic binding between Dsc2apATH andDsg2pATH monomers were ,0.23 s21 and that they werestatistically indistinguishable from each other. These results

indicate that monomeric Dsc2a, but not Dsg2, forms Ca2+-dependent homophilic interactions, and that Dsc2a forms Ca2+-independent heterophilic interactions with Dsg2.

W2 is required for Dsc2a Ca2+-dependent homophilic bindingand recruitment into desmosome punctaThe amino acid sequence of the extracellular domain of classical

and desmosomal cadherins contains a tryptophan at position 2

Fig. 1. Localization of desmoplakin I/II to surfacecomplexes depends on desmosomal cadherinengagement. MDCK cells were plated at single cell densityonto micro-patterned substrates functionalized with collagenIV and either Fc (A), Dsg2Fc (B) or Dsc2aFc (C), or Dsg2Fcplus Dsc2aFc (D). After 4–6 h, cells were fixed andimmunostained for the desmosomal adaptor proteindesmoplakin (DP I/II). Localization of DP I/II at the cell–substrate interface, and collagen IV stripe location (Cy3),were visualized using TIRF microscopy. Most cells showedthis behavior (n.5) and a representative cell was chosen foreach condition; merged images show specific recruitment ofDP I/II with respect to stripe functionalization and location.Dsc2aFc, Dsg2Fc and Fc control stripes are marked as thenon-collagen IV printed surfaces (black stripes). The whiteline designates the cell outline. Scale bar: 10 mm.

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(W2) in the first extracellular domain (EC1). W2 in classicalcadherins is crucial for trans binding between opposingextracellular domains (Shapiro et al., 1995) because it forms a

strand-swap dimer between opposing cadherins (Boggon et al.,2002). A crystal structure of desmosomal cadherins has not beenreported, but it has been suggested that desmosomal cadherins use

a similar binding mechanism (Nie et al., 2011). To test for a role ofW2, we made alanine substitution mutants in Dsc2a (Dsc2aW2A)and Dsg2 (Dsg2W2A), and monomeric extracellular domain

mutants containing the pATH tag (Dsc2aW2ApATH,Dsg2W2ApATH) were used for SMFS (Fig. 3A).

The W2A mutation resulted in loss of Ca2+-dependent

Dsc2apATH homophilic binding measured by SMFS, becausethe cadherin binding frequency did not change when free Ca2+

was chelated by EGTA (Fig. 3B), unlike the wild-type

Dsc2apATH (see Fig. 2E). However, the frequency of Ca2+

independent heterophilic binding between Dsc2aW2ApATH andDsg2W2ApATH (Fig. 3B) was similar to the wild-type protein

binding (Fig. 2E). These results indicate that homophilic Dsc2a–Dsc2a binding is Ca2+ dependent, and requires a strand-swapmechanism involving W2. In contrast, heterophilic binding

between Dsc2a and Dsg2 is Ca2+ independent and occursthrough a W2-independent mechanism. This is in contrast to E-cadherin, in which mutations of both W2 and K14 residues are

required to inhibit binding (Rakshit et al., 2012).We tested whether the Dsg2W2A and Dsc2aW2A mutants

could be incorporated into endogenous desmosome puncta in

contacting MDCK cells. C-terminal GFP-tagged mutant proteinwas transiently expressed in MDCK cells and their distributions(linear versus punctate) were compared to transiently expressed

Fig. 2. Desmosomal cadherins havedistinct extracellular trans bindingproperties. (A) A schematic of themonomeric desmosomal cadherinsfused to tags comprising Avadin, Tevand His (pATH). (B). Coomassie-bluestained SDS-PAGE gel of purifiedDsg2pATH, Dsc2apATH and E-cadherinpATH (EcadpATH, control),which have similar molecular masses(,95 kDa); an unprocessed precursorform of the cadherins is present at lowlevels in the Dsc2apATH sample (*).(C) Schematic of an AFM cantilever tipand coverslip coated with PEG (5%coverage), and functionalized withmonomeric Dsg2pATH or Dsc2apATHto measure single-moleculeinteractions. (D) The AFM cantilever tipwas lowered and raised thousands oftimes to measure hundreds of single-molecule interactions between proteinon the AFM tip and protein on thecoverslip, and only events showing asingle binding event were used foranalysis. (E) Single-molecule bindingfrequency of different combinations ofDsg2pATH and Dsc2apATH wasplotted as a percentage of all eventsand compared to non-specific binding(identically functionalized cantileverslacking the desmosomal cadherins).Addition of the Ca2+ chelator EGTAwas used to test Ca2+-dependence ofbinding events. (F) The relationshipbetween loading rate (pN/s) and bondrupture force was plotted for bothhomophilic binding of Dsg2pATH (withor without EGTA) and Dsg2pATH–Dsc2apATH heterophilic binding.

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wild-type protein and/or endogenous desmosomal proteins

(Fig. 4). Dsg2W2A–GFP was restricted to puncta along cell–cellcontacts, similar to wild-type Dsg2–GFP (Fig. 4A). HoweverDsc2aW2A–GFP had a linear distribution along the plasma

membrane at cell-cell contacts (Fig. 4B, C) compared to wild-type Dsc2a–GFP (Fig. 4D,E). Mislocalization of Dsc2aW2A–GFPwas not due to overexpression since the transfection efficiency andexpression levels were similar to control Dsc2a–GFP levels

(Fig. 4F).Next we investigated whether the anomalous localization of

Dsc2aW2A–GFP along cell–cell contacts affected the recruitment

and localization of endogenous components into desmosomepuncta. MDCK cells transiently co-expressing Dsc2aW2A–GFPand Dsc2a–DsRED (Fig. 4B,C), or only Dsc2a–GFP (Fig. 4D,E)

were fixed and stained for the desmosome proteins DP I/II, thecytokeratin 8 and cytokeratin 18 pair (hereafter cytokeratin 8/18)(Fig. 4B,D) or plakoglobin (Fig. 4C,E). A representative cell–cell

contact with an expressing cell is shown for each staining. Line-scans that measured fluorescence intensity for each protein werenormalized and used to determine whether endogenous proteinscolocalized with the wild-type or W2A mutant Dsc2a, and

whether the localization had a linear or punctate distribution(Fig. 4B–E).

In contrast to the linear plasma membrane staining of

Dsc2aW2A–GFP, endogenous DP I/II, cytokeratin 8/18, wild-type Dsc2a–DsRED and wild-type Dsc2a–GFP all localized inpuncta along the same cell–cell contacts (Fig. 4B,D).

Endogenous plakoglobin staining was also punctate, and moresimilar to wild-type Dsc2a–DsRED and Dsc2a–GFP than mutantDsc2aW2A–GFP (Fig. 4C,E). These results indicate that bothendogenous desmosomal cytoplasmic proteins and cadherins

localized normally in puncta at cell–cell contacts in the presenceof Dsc2aW2A, suggesting that Dsc2aW2A did not act as adisrupting (dominant-negative) protein and that it was mostly

excluded from endogenous desmosome puncta.

Dsc2aW2A mutant is more mobile than wild-type Dsc2a atcell–cell contactsWe speculated that the lack of incorporation into endogenousdesmosome puncta would result in a higher mobility of

Dsc2aW2A in the membrane than Dsc2a, Dsg2 and Dsg2W2A,all of which were incorporated into desmosomal puncta.Fluorescence recovery after photobleaching (FRAP) was used

to measure the mobile fraction and recovery rate of wild-type and

mutant proteins based on a single exponential recovery model(Fig. 5). Dsg2W2A had a slightly higher mobile fraction (0.354;P,0.1) and slower recovery rate (t1/25167 s; P,0.05) than

wild-type Dsg2 (0.312 and 127 s, respectively) (Fig. 5B;supplementary material Movies 1, 2). Although the recoveryrate of Dsc2aW2A was similar to that of Dsg2W2A (174 versus167 s, respectively; Fig. 5B; supplementary material Movie 3),

the Dsc2aW2A mobile fraction was significantly greater(P,0.0001) than that of Dsc2a (0.714 versus 0.351; Fig. 5B;supplementary material Movie 4). Indeed the mobile fraction of

Dsc2aW2A was similar to that of Lyn–GFP, a lipid-anchoredprotein (0.714 and 0.75, respectively; Fig. 5B).

These FRAP data indicate that the recruitment of wild-type

Dsc2a and Dsg2 into desmosome puncta results in a reducedmobile fraction that might be due to their incorporation into adense protein complex with adaptor proteins and intermediate

filaments (see Fig. 4). The Dsg2W2A mutant had a reducedmobile fraction indicating that W2 and strand-swap dimers arenot required for Dsg2 incorporation into desmosome puncta. Incontrast, the Dsc2aW2A mutant was distributed along the plasma

membrane and had a mobile fraction similar to a lipid-anchoredprotein, confirming that W2 and strand-swap dimers are requiredfor Dsc2a incorporation into desmosome puncta and that the

W2A mutant was likely either not incorporated or retained inthese structures.

Desmosome formation inhibits MDCK cell migrationIn addition to testing protein recruitment to different adhesioncomplexes, dual patterned substrates test the effects of cell–celladhesion proteins on cell migration on ECM (Borghi et al., 2010).

Although E-cadherin had little effect on the rate of cell migrationon collagen IV (Borghi et al., 2010), we found that adhesion todesmosomal cadherins had a significant effect. To quantify cell

movements, images were recorded for 6–10 h, and changes innucleus position were tracked to determine the migration velocity(Fig. 6A) and migration direction (Fig. 6B). In addition, the

contour of a representative cell was traced over time to examinelamellipodia dynamics (Fig. 6C).

Migration velocities of MDCK cells on all surfaces were

normalized to 10 mm/h and plotted on a natural log (ln) scale forcomparison. The migration rate of cells plated on collagen IV andDsg2Fc substrate stripes (,9 mm/h) was similar to that of cells

Fig. 3. A W2A mutation in Dsc2a disruptssingle-molecule trans binding. (A) Atryptophan residue at position 2 (W2) in thefirst extracellular binding repeat (EC1) wasmutated to an alanine residue (W2A) inDsg2W2ApATH and Dsc2aW2ApATH.Coomassie-stained SDS-PAGE shows amajor protein band of the expected molecularmass, and low-level contamination bypossible dimers (*; ,200 kDa) anddegradation products (**). A lane between thesamples has been deleted for simplificationpurposes (dashed line). (B) SMFS showedlow levels of Ca2+-independent homophilicbinding of Dsg2W2ApATH andDsc2aW2ApATH . Ca2+-independentheterophilic binding of Dsg2W2ApATH andDsc2aW2ApATH was comparable to that inwild type (Fig. 2E).

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plated on collagen IV and Fc control substrate stripes (Borghi

et al., 2010), and was significantly faster than cells on collagen IVand Dsc2aFc substrate stripes (,5 mm/h; P,0.01). Importantly,the migration rate of cells plated on collagen IV and Dsg2Fc plus

Dsc2aFc substrate stripes was in between that of cells on collagenIV and Dsc2aFc stripes or collagen IV and Dsg2Fc stripes(,7 mm/h; P,0.1).

The directionality of cell movements on the dual-patterned

surfaces were plotted as cell velocity in the direction of the stripesdivided by the perpendicular velocity (x/y); ln(x/y) is positive ifcells preferentially move in the direction of the stripes (x.y). Cell

migration on collagen IV and Dsg2Fc stripes showed biasedmovement in the direction of the stripes (Fig. 6B), as reportedpreviously for cells migrating on collagen IV and E-cadherinFc

Fig. 4. See next page for legend.

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stripes (Borghi et al., 2010). Directional bias is due to restrictionof the migration-based adhesion machinery, such as vinculin–GFP[supplementary material Fig. S3 (see also Borghi et al., 2010)],

to the collagen IV stripes. In contrast, cells were almoststationary on collagen IV and Dsc2aFc stripes and the smallamount of movement only showed a slightly significant

difference from Dsg2Fc (Fig. 6B; P,0.1). The direction ofMDCK cell migration on collagen IV and Dsg2Fc plus Dsc2aFcstripes was significantly biased towards the direction of the

stripes (P,0.0001; supplementary material Movie 5) comparedto Dsc2aFc alone and slightly significant compared to Dsg2Fcalone (P,0.1).

We examined membrane lamellipodia activity by tracing thecell contour over time and superimposing all images onto thedual-patterned substrate (Fig. 6C); quantification of membranedynamics was not attempted because the cells migrated during the

course of experiment. Cells on collagen IV and Dsg2Fc stripesexhibited membrane dynamics over the whole cell contour, butpersistent membrane projections occurred predominantly in the

direction of cell movement, which was generally along thecollagen IV stripes (Fig. 6C; supplementary material Movie 6).Cells on collagen IV and Dsc2aFc stripes exhibited membrane

dynamics, but few persistent membrane projections (Fig. 6C;supplementary material Movie 7) consistent with the significantreduction in the overall migration rate (Fig. 6A). Cells oncollagen IV and Dsg2Fc plus Dsc2aFc stripes had membrane

dynamics similar to cells on Dsg2Fc, and more persistentmembrane projections on the collagen IV stripes, which wasaccompanied by highly processive cell movement (Fig. 6C;

supplementary material Movie 5) consistent with a greaterdirectional migration bias than on collagen IV and Dsg2Fcstripes, collagen IV and Dsc2aFc stripes (Fig. 6B; Movies 6, 7) or

collagen IV and Fc control substrates (Borghi et al., 2010).These results indicate that desmosome puncta formed by

homophilic Dsc2a binding (see Fig. 1) result in strong cell

adhesion to the Dsc2aFc stripe. We suggest that the smallervelocity decrease in the presence of Dsc2aFc plus Dsg2Fc wasdue to the dilution of Dsc2aFc homophilic interactions by

Dsg2Fc, which results in decreased adhesion strength and therebyincreased cell migration. Thus, migration direction bias requires

both anisotropic adhesion of integrin-based complexes on thecollagen IV substrate and reduced adhesion strength fromdesmosome assembly due to the incorporation of Dsg2.

DISCUSSIONDesmosomes assemble in a Ca2+-dependent manner betweenopposing cells, and are required to maintain the structuralintegrity of tissues. Dsg and Dsc are obligate cadherins in

desmosomes, and impairment of either Dsc or Dsg extracellularinteractions causes heart defects, and epidermal blisteringdiseases and syndromes (Al-Jassar et al., 2013; Kottke et al.,

2006; Thomason et al., 2010). Studies using different in vitro

binding and cell-based assays, however, have not identifiedspecific roles for Dsg and Dsc in desmosome assembly andadhesion. Whether there is specificity in the extracellular binding

properties of desmosomal cadherins has been difficult to establishowing to the overlapping roles of other cadherins (e.g. E-cadherin) in epithelial cell–cell adhesion, the complexity of

proteins in desmosomes and their resistance to dissociation innon-denaturing conditions (Nie et al., 2011), and the lack of high-resolution crystal structures of the two desmosomal cadherin sub-

types (Al-Jassar et al., 2013). Here, we used reductionistapproaches to distinguish roles of Dsc2a and Dsg2 indesmosome assembly, organization and adhesion in the absenceof other cell–cell adhesions. Our results indicate that Dsc2a and

Dsg2 have distinct binding properties and functions indesmosome organization and adhesion.

Micro-patterned substrates of purified Dsc2aFc or Dsg2Fc

revealed that Dsc2aFc, but not Dsg2Fc, was necessary andsufficient to induce the recruitment of a desmosome-specificcytoplasmic protein (e.g. DP I/II) into punctate cellular structures

(Fig. 1). Given that MDCK cells express both Dsc2a and Dsg2(Pasdar and Nelson, 1989; Pasdar et al., 1991), the exogenoussubstrate-bound Dsc2a could have initiated the assembly of

desmosome puncta by binding cellular Dsc2a, Dsg2 or both onthe ventral surface of the cell. Indeed, our results from the SMFSassay indicated that Dsc2a forms homophilic interactions andheterophilic interactions with Dsg2 (Fig. 2). However, SMFS

data indicates only heterotypic interactions between Dsg2 andDsc2a, suggesting that Dsg2Fc surfaces promote heterotypicbinding and diffuse recruitment of DP I/II at the membrane.

Interestingly, other studies using chemical cross-linkers onlyfound homophilic binding by Dsc2 and Dsg2 (Nie et al., 2011),whereas bulk biochemical assays indicate heterophilic Dsc2a–

Dsg2 binding and weak homophilic Dsg2–Dsg2 binding inagreement with our SMFS results (Syed et al., 2002). Differencesbetween other SMFS results (Hartlieb et al., 2013; Schlegel et al.,2010) and our SFMS experiments might be due to our strict

definition of a single-molecule interaction or our use ofmonomeric proteins, which excluded the possibility of inducedlateral (cis) clustering of extracellular domains. In addition, our

findings of differences in the Ca2+ dependency of homophilic andheterophilic protein binding have revealed differences betweensingle-molecule data and bulk or cellular assays (see below).

Analysis of the crystal structure of classical cadherins revealedthat a tryptophan at position 2 (W2) in the N-terminal EC1domain is crucial for the formation of a strand-swap dimer

between opposed extracellular domains (Boggon et al., 2002); anintermediate adhesion structure is also formed by a weak X-dimerstructure (Harrison et al., 2010). Although the atomic structures

Fig. 4. The W2A mutation disrupts localization of Dsc2a to punctatecomplexes. (A) MDCK cells were transiently transfected with wild-type GFP-tagged Dsg2 or GFP-tagged Dsg2W2A and fixed with methanol. Wild-typeDsg2–GFP and mutant Dsg2W2A–GFP localized in characteristicdesmosome-like punctate spots at cell–cell contacts. Scale bar: 10 mm.(B–E) MDCK cells were transiently transfected with GFP-tagged Dsc2aW2Aor wild-type GFP-tagged Dsc2a, with or without RFP-tagged wild-typeDsc2a, fixed and stained for endogenous desmosomal proteins. Top: a wide-field image with the representative contact outlined in white. Middle: zoomed-in images of the representative cell–cell contacts. Bottom: line scans ofnormalized fluorescence intensity for Dsc2aW2A–GFP, Dsc2a–GFP, Dsc2a–DsRED, plakoglobin (PG) and DP I/II were plotted for the representative cell–cell contact. Intensity was normalized by subtracting the average intensityover the whole contact and dividing by the difference in maximum andminimum values (pixel range). (B,C) Dsc2aW2A–GFP accumulated in alinear, non-punctate distribution along the plasma membrane and did not co-localize with desmosomal proteins (DP I/II, cytokeratin 8/18, and plakoglobin)or co-transfected wild-type Dsc2a_DsRED, all of which had acharacteristic desmosome-like punctate distribution (D,E) Wild-type Dsc2a–GFP localized into punctate spots at cell–cell contacts and colocalized withthe desmosomal proteins (DP I/II, cytokeratin 8/18 and plakoglobin). Of note,the focal plane was set using the staining of endogenous proteins as themarker, so as not to create any biases between the transfected GFP-taggedproteins. Scale bar: 5 mm. (F) Expression levels of transiently transfectedcells (similar efficiencies with .60% transfected) Dsc2aW2A–GFP were ,1/2 that of endogenous Dsc2a in MDCK cells.

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of full-length Dsc2a or Dsg2 have not been solved, both proteinshave a tryptophan at position 2 (Nie et al., 2011) but neither havethe necessary residues for the formation of an X-dimer.Significantly, the W2A mutation in Dsc2a inhibited Ca2+-

dependent homophilic binding (Fig. 3). Furthermore, theDsc2aW2A mutant appeared to be excluded from endogenousdesmosomes in MDCK cells, which we inferred from its linear

plasma membrane staining compared to endogenous desmosomepuncta in the same cells (Fig. 4), and significantly increasedmobile fraction compared to wild-type Dsc2a (Fig. 5). Given that

Dsc2aFc alone was necessary and sufficient to inducedesmosome-like DP I/II puncta in cells on micro-patternedsubstrates (Fig. 1), we suggest that Dsc2a homophilic bindingthrough a Ca2+- and W2- (strand-swap dimer) dependent

mechanism is required for desmosome assembly. Furthermore,because Dsc2aW2A was mostly excluded from desmosomes anddid not affect the punctate localization of desmosome adaptor

proteins (Fig. 4), it is likely that the W2 strand-swap mechanismis also necessary for the incorporation of Dsc2a into desmosomepuncta, and that binding to cytoplasmic adaptor proteins is not

sufficient.In contrast, the W2A mutation in Dsg2 affected neither Ca2+-

independent heterophilic binding to Dsc2a (Fig. 3), colocalization

with endogenous desmosome puncta in MDCK cells (Fig. 4) norits mobile fraction (Fig. 5). These results indicate that Dsg2incorporation into desmosomes occurs through a Ca2+- and

W2- (strand-swap dimer) independent mechanism that relies onheterophilic interactions and/or cytoplasmic interactions withother proteins in the desmosome. Given that Dsg2Fc alone did notinduce assembly of desmosomal-like DP I/II puncta in MDCK

cells on micro-patterned substrates (Fig. 1), what role couldDsg2-dependent Ca2+-independent heterophilic adhesion play?Differences in Ca2+-dependency of desmosomal adhesion have

been identified in tissue- and cell-based studies, and it has beensuggested that mature desmosome complexes are in a hyper-adhesive state that is Ca2+ independent (Garrod and Kimura,

2008). Our results raise the possibility that such Ca2+-independentadhesion might be mediated by heterophilic Dsc2a–Dsg2 binding.

To discriminate the effects of desmosome adhesion on cellmigration from the contribution of other adhesion mechanisms

(e.g. classical cadherins), we examined cell migration on dual-patterned substrates comprising alternating stripes of collagen IVand Dsc2aFc and Dsg2Fc (Figs 1, 2). We can infer the

contribution of desmosome assembly on cell adhesion bymeasuring the rate of cell migration. Significantly, MDCK cellmigration was severely inhibited on dual-patterned substrates

coated with saturating amounts of Dsc2aFc (Fig. 6A;supplementary material Movie 7); in contrast, Dsg2Fcsubstrates did not inhibit cell migration compared to controls

(Fig. 6A; supplementary material Movie 6). Our previous studieswith substrates coated with E-cadherinFc and collagen IVrevealed little or no effect of E-cadherin on the migration rate

Fig. 5. Membrane mobility of Dsc2aincreases in the W2A mutant. (A) FRAPof GFP-tagged wild-type and W2A mutantDsc2a and Dsg2 cadherins at cell–cellcontacts showed a significant difference inthe recovery rate of Dsc2aW2A and wild-type Dsc2 and Dsg2, whereas mutantDsg2W2A showed only slight differences.Lyn–GFP is used a reference for proteinsthat are lipid-anchored to the plasmamembrane. Plotted as mean6s.d.(B) Mobile factions and recovery rates (t1/2)are compared using a single exponentialcurve fitted to the data shown in A. Mobilefractions of Dsc2a–GFP and Dsc2aW2A–GFP are significantly different (P,0.0001),whereas Dsg2–GFP and Dsg2W2A–GFPare less significant (P,0.1). The recoveryrates of both pairs are significantly different(P,0.05). Significance was calculatedusing a comparison of fits, extra sum-of-squares F test.

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of MDCK cells (Borghi et al., 2010). Therefore, we propose that theformation of desmosome-like structures, unlike E-cadherinadhesion, mechanically resists cell movement by forming strongadhesive bonds to the Dsc2aFc substrate, rather than a

downregulation of the Arp2/3 and actomyosin machineries thatare involved in membrane dynamics and cell migration (Pollard andBorisy, 2003). Indeed, whereas Dsc2aFc-containing dual substrates

inhibited cell migration, we found that they had little effect onoverall lamellipodia dynamics (Fig. 6C). Interestingly, addition ofDsg2Fc to the Dsc2aFc substrate partially rescued the reduction in

cell migration rate (Figs 6A,C). We suggest that differences in theratio of Dsc2a and Dsg2 in desmosomes might regulate the strengthof cell–cell adhesion and dynamics of the desmosome complex

thereby modulating cell migration in epithelial sheets. However,further studies are needed to test this hypothesis in detail.

In summary, our results have uncovered a specific role forextracellular contacts formed by Dsc2a in the structural

organization and function of desmosomes. The assembly ofdesmosome puncta depends on Ca2+- and W2- (strand-swapdimer) dependent homophilic trans-dimerization between Dsc2a

proteins on opposing cell surfaces. Dsg2 might be required for thelong-term stability of desmosomes, and also perhaps theformation of a Ca2+-independent hyperadhesive state (Garrod

and Kimura, 2008). Whereas Dsc2a might have a specific role in

adhesion, Dsg2 might have additional roles (Brennan andMahoney, 2009; Brennan et al., 2007). Thus, differences in themechanisms of incorporation and function of Dsc and Dsgcadherins might allow a diversification of desmosome functions

in cell adhesion, migration and differentiation.

MATERIALS AND METHODSProtein purificationThe cloning design for desmosomal Fc fusion proteins was described

previously for E-cadherinFc (Drees et al., 2005). The C-terminal of the

Dsg2 or Dsc2a extracellular domains was fused to the constant region of

human IgG1 (Fc). Fusion constructs were expressed in HEK293T cells and

proteins purified from conditioned medium over Protein A Sepharose (GE

Healthcare Life Sciences, Uppsala, Sweden). Proteins were then labeled

with Cy5.5 dye (Invitrogen Life Technologies, Carlsbad, CA, USA) for

visualization. Monomeric Dsg2 and Dsc2a construct design was also as

described previously (Zhang et al., 2009), where the C-terminal of the

extracellular domain was fused to the Avi tag, Tev sequence and His tag

(pATH). Fusion constructs were expressed in HEK293T cells, and proteins

were purified from conditioned medium over Ni-NTA agarose beads, and

protein was biotinylated using BirA enzyme for surface functionalization

(BirA500 kit, Avidity LLC, Aurora, CO, USA).

Surface immobilization of desmosomal cadherinsDual-micropatterned surfaces were prepared using a two-step process as

reported previously (Borghi et al., 2010). First, the collagen IV was

Fig. 6. Desmosomal cadherin engagement regulates MDCK cell migration on dual-patterned substrates. The movement of single MDCK cells on eachdual-patterned substrate condition was tracked by monitoring changes in nuclear position over time. (A) Cell migration rate (v) was normalized to single cellmovement on only collagen IV surfaces, and plotted as a tukey box-and-whisker plot on a natural log scale [Ln(v/10 mm/h)]. The box marks the range fromthe 25th–75th percentile of the data and the band inside the box marks the median. Whiskers show the maximum and minimum values. (B) Directionality ofmovement was plotted as a tukey box-and -whisker plot on a natural log scale as a function of movement parallel to stripes (x) divided by perpendicularmovement (y). *P,0.1; **P,0.01; ****P,0.0001. (C) Membrane activity during migration assays was assessed over time by tracing the cell contour from DICimages and projecting the outlines onto the stripe patterns (C). Statistical analysis used the Mann–Whitney, two-tailed t-test (Dsg2Fc, n513; Dsc2aFc, n531;Dsg2Fc+Dsc2aFc, n527).

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micro-contact printed using standard protocols (Cuvelier et al., 2003;

Nishizawa et al., 2002), and then the non-ECM–coated surfaces were

functionalized with by fusion with the human IgG1 Dsg2aFc, Dsc2aFc or

Dsg2aFc plus Dsc2aFc as described previously (Drees et al., 2005).

Polydimethylsiloxane (PDMS) stamps with striped groves were coated

with Cy3.5-tagged collagen IV (200 mg/ml) and printed onto silanized

coverslips. Stamped coverslips were first incubated with EZ link sulfo-

NHS-LC-LC-biotin (50 mM in water; made fresh each time), then

neutravidin (5 mg/l in PBS), then biotinylated Protein A (0.3 mg/ml in

PBS), each for 1 h. Before incubation with Dsc2aFc or Dsg2aFc protein,

coverslips were blocked with a D-biotin solution (15 mM in DMSO) for

30 min. Coverslips were washed with PBS three times between

incubations and all incubations and washes were performed at room

temperature in the dark. Finally, coverslips were incubated with Cy5.5-

tagged Dsg2aFc, Dsc2aFc or Fc protein (control) for 1 h at room

temperature. MDCK cells were added to the coverslips at low density.

For single-molecule force spectroscopy (SMFS), AFM cantilevers and

glass coverslips were first cleaned with a 25% H2O2:75% H2SO4 solution

and subsequently washed with deionized water, 1 M potassium hydroxide

solution, deionized water and acetone. The cleaned AFM cantilevers and

coverslips were made amine reactive by functionalizing with 2% v/v

solution of 3-aminopropyltriethoxysilane (Sigma-Aldrich, St. Louis, MO,

USA) dissolved in acetone. The cantilevers and coverslips were then

functionalized with PEG spacers (PEG 5000, Laysan Bio, Arab, AL,

USA) containing an amine-reactive N-hydroxysuccinimide ester at one

end to bind to the surface. A known fraction of PEG presented biotin at

the other end and was kept low in order to measure single-molecule

events; for wild-type and mutant monomers, 5% and 7% of the PEG

contained biotin, respectively. The biotinylated surfaces were incubated

with 0.1 mg/ml BSA for 12 h, to minimize non-specific surface

interactions, and then with 0.1 mg/ml streptavidin for 30 min.

Biotinylated Dsc2a or Dsg2a monomers were bound to streptavidin on

the AFM tip and coverslip surfaces. Following Dsc2a or Dsg2a

immobilization, the surfaces were incubated in 10 mM biotin to block

free biotin-binding sites on streptavidin.

Single molecule dynamic force spectroscopy experimentsSingle molecule dynamic force spectroscopy experiments were

performed at different pulling velocities using an Agilent 5500 AFM.

Experiments were performed in 10 mM Tris-HCl pH 7.5, 100 mM NaCl,

10 mM KCl in either 2.5 mM CaCl2 or 2 mM EGTA. Single unbinding

force curves were selected and fitted with the freely jointed chain (FJC)

model using the following equation (Smith et al., 1992):

l Fð Þ~Lc cothFa

kBT

� �{

kBT

Fa

� �, ð1Þ

where l Fð Þ is the end-to-end distance of the PEG tether subjected to a

force F, Lc, is the contour length of PEG and a is the Kuhn length. Spring

constants of the AFM cantilevers were measured with the thermal

fluctuation method (Hutter and Bechhoefer, 1993). In order to measure

unbinding forces at different loading rates, the cantilever was mounted on

a piezoelectric translator and was moved at different velocities (v); the

loading rate (vF) was estimated using (Marshall et al., 2006; Ray et al.,

2007) the equation:

1

vF

~1

kcvz

1

v

dl Fð ÞdF

~1

kcv1z

kcLca

kBT

kBT

Fa

� �2

{csch2 Fa

kBT

� � !" #, ð2Þ

where kc is the spring constant of the cantilever.

In order to eliminate hidden multiple unbinding events from visibly

indistinguishable single unbinding events, we followed a protocol used

for identifying the heterogeneity in chemical bonds or hidden multiple

bonds as described previously (Fuhrmann et al., 2012). After eliminating

all hidden multiple events, the forces were plotted as a histogram for each

retracting velocity with bin width calculated according to a statistical

method (Scott, 1979). The histograms were fitted to a Gaussian

distribution to obtain the most probable forces (Fmp) for each retracting

velocity. Most probable force (Fmp) vs mean loading rate (vF ) data were

then fitted (Bell, 1978; Evans and Ritchie, 1997) in order to measure the

force-independent off-rate (k0off ) and the distance (xb) from the minimum

of the reaction potential to the peak of the potential-barrier using the

equation:

Fmp~kBT

xbln

vFxb

k0off kBT

!: ð3Þ

The statistical significance of parameters obtained from the fits was

tested using a Student’s t-test.

Cell culture and constructsMadin–Darby canine kidney (MDCK) type II G cells were grown in

DMEM with low glucose and 200 mM G418 for stably expressing tagged

proteins of interest (Vin–GFP, E-cadherin–DsRED). Cells stably or

transiently expressing fluorescently tagged proteins were monitored on a

widefield epifluorescence inverted microscope equipped with TIRF-M

and FRAP lasers. Desmosomal cadherin mutants were generated from a

Dsg2–GFP construct and a Dsc2a–GFP construct (gift from the

laboratory of Sergey M. Troyanovsky, Feinberg School of Medicine,

Northwestern University, IL) using site-directed mutagenesis (Aligent

Technologies, Stratagene, Santa Clara, CA, USA) with PCR fragments

containing the point mutation for monomeric fusion proteins

(Dsg2aW2A_pATH, Dsc2aW2A_pATH) and full-length GFP-tagged

versions (Dsg2aW2A–GFP and Dsc2aW2A–GFP).

Fluorescence microscopyImmunofluorescence was performed on MDCK cells fixed with 4% PFA

for 20 min at 25 C or 100% methanol for 5 min at 220 C, and extracted

with 0.1% Triton X-100 for 5 min before blocking in PBS containing

0.2% BSA, 50 mM NH4Cl2 and 1% normal goat/donkey serum. Cells

were incubated with antibodies against the proteins of interest: anti-

desmoplakin (DP I/II) (Pasdar and Nelson, 1988a), anti-plakoglobin

(clone 15F11 – Thermo Fisher Scientific, Waltham, MA, USA), anti-

desmocollin2 (Dsc2a) (ab72792 – Abcam, Cambridge, England), anti-

cytokeratin 8/18 [Abcam (Cy90) – ab17151]. Cells were incubated with

secondary (Fab9) goat/donkey anti-mouse/rabbit IgG labeled with FITC/

Rhodamine Red-X/Cy5 (1:200; Jackson ImmunoResearch Laboratories,

Westgrove, PA, USA) and mounted using Vectashield (Vector Labs,

Burlingame, CA, USA) (or PBS for TIRF imaging). Immunofluorescence

images were acquired on a Zeiss (Jena, Germany) Axiovert 200 inverted

microscope equipped with a Mercury lamp and a 1006 objective

(Olympus, Tokyo, Japan) and acquired with AxioVision Microscopy

Software (Zeiss). Fluorescence live-cell imaging (FRAP) and TIRF

images were acquired on a custom-built Zeiss Axiovert 200M inverted

widefield epifluorescence microscope [Intelligent Imaging Innovations

(3i), Denver, CO, USA] equipped with including a 175 Watt Xenon light

source with a dual galvanometric filter changer, a Coolsnap HQ interline

CCD camera (Photometrics, Tuscon, AZ, USA), an x,y motorized stage

with harmonic drive z-focusing, a quad filter set for DAPI, FITC, Cy3

and Cy5, and a MicroPoint FRAP laser system (Andor Technology,

South Widsor, CT, USA), as described previously (Spiliotis et al., 2005;

Yamada et al., 2005). Live cells were imaged in DMEM without Phenol

Red, supplemented with 25 mM Hepes at 37 C on a 1006 1.4 NA oil

objective (FRAP and TIRF) and a 406air objective for tracking cellular

movement; images were acquired using Slidebook (3i) software. The

fluorescence signal from cells (GFP) and micro-patterned substrates

(Cy3.5 or Cy5.5 dye) was used to assess protein recruitment and cell

position at long time scales for measurement of cell migration, whereas

DIC images at long time scales were used to monitor cell contour outlines

and fluctuations. Image analysis was performed with Image J and

statistical analysis was performed using a Mann–Whitney, two-tailed t

test. FRAP data was normalized and fitted with a single exponential

recovery model as described previously (Borghi et al., 2010). Significant

differences between modeled parameters were calculated using a

comparison of fits, extra sum-of-squares F test.

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Competing interestsThe authors declare no competing interests.

Author contributionsM.L. designed, performed and interpreted the experiments, assembled thefigures, and wrote the manuscript; S.R., O.S. and S.S. performed the SMSFexperiments; R.M.H. provided the Dsg-Fc and Dsc2-Fc fusion proteins; N.B. wasinvolved in the initial design of experiments; K.J.G., S.S. and W.J.N. were involvedin designing the experiments and writing the manuscript.

FundingWork in the S.S. laboratory was partially supported by a grant from the AmericanHeart Association (Scientist Development Grant 12SDG9320022). Work in theK.J.G. laboratory was supported by grants from the National Institutes of Health(NIH) [grant numbers R37AR043380, R01 AR41836]. Work in the W.J.N.laboratory was supported by the NIH [grant number GM35527] and HumanFrontier Science Program [grant number RGP0040/2012]. M.L. was supported bya Genentech Graduate Fellowship, Cancer Biology Training Grant and MasonCase Graduate Fellowship. R.M.H. was supported by a NIH T32 training grant[grant number GM08061]; and an American Heart Association predoctoralfellowship. Deposited in PMC for release after 12 months.

Supplementary materialSupplementary material available online athttp://jcs.biologists.org/lookup/suppl/doi:10.1242/jcs.146316/-/DC1

ReferencesAl-Amoudi, A., Dıez, D. C., Betts, M. J. and Frangakis, A. S. (2007). Themolecular architecture of cadherins in native epidermal desmosomes. Nature450, 832-837.

Al-Amoudi, A., Castano-Diez, D., Devos, D. P., Russell, R. B., Johnson, G. T.and Frangakis, A. S. (2011). The three-dimensional molecular structure of thedesmosomal plaque. Proc. Natl. Acad. Sci. USA 108, 6480-6485.

Al-Jassar, C., Bikker, H., Overduin, M. and Chidgey, M. (2013). Mechanisticbasis of desmosome-targeted diseases. J. Mol. Biol. 425, 4006-4022.

Amagai, M. and Stanley, J. R. (2012). Desmoglein as a target in skin disease andbeyond. J. Invest. Dermatol. 132, 776-784.

Bell, G. I. (1978). Models for the specific adhesion of cells to cells. Science 200,618-627.

Bhuiyan, Z. A., Jongbloed, J. D. H., van der Smagt, J., Lombardi, P. M.,Wiesfeld, A. C. P., Nelen, M., Schouten, M., Jongbloed, R., Cox, M. G. P. J.,van Wolferen, M. et al. (2009). Desmosome mutations in arrhythmogenic rightventricular cardiomyopathy: important insight but only part of the picture. Circ.Cardiovasc. Genet. 2, 415-417.

Boggon, T. J., Murray, J., Chappuis-Flament, S., Wong, E., Gumbiner, B. M.and Shapiro, L. (2002). C-cadherin ectodomain structure and implications forcell adhesion mechanisms. Science 296, 1308-1313.

Borghi, N., Lowndes, M., Maruthamuthu, V., Gardel, M. L. and Nelson, W. J.(2010). Regulation of cell motile behavior by crosstalk between cadherin- andintegrin-mediated adhesions. Proc. Natl. Acad. Sci. USA 107, 13324-13329.

Bornslaeger, E. A., Corcoran, C. M., Stappenbeck, T. S. and Green, K. J.(1996). Breaking the connection: displacement of the desmosomal plaqueprotein desmoplakin from cell-cell interfaces disrupts anchorage of intermediatefilament bundles and alters intercellular junction assembly. J. Cell Biol. 134, 985-1001.

Brennan, D. and Mahoney, M. G. (2009). Increased expression of Dsg2 inmalignant skin carcinomas: A tissue-microarray based study. Cell Adhes. Migr.3, 148-154.

Brennan, D., Hu, Y., Joubeh, S., Choi, Y. W., Whitaker-Menezes, D., O’Brien,T., Uitto, J., Rodeck, U. and Mahoney, M. G. (2007). Suprabasal Dsg2expression in transgenic mouse skin confers a hyperproliferative and apoptosis-resistant phenotype to keratinocytes. J. Cell Sci. 120, 758-771.

Chidgey, M. and Dawson, C. (2007). Desmosomes: a role in cancer? Br.J. Cancer 96, 1783-1787.

Chitaev, N. A. and Troyanovsky, S. M. (1997). Direct Ca2+-dependentheterophilic interaction between desmosomal cadherins, desmoglein anddesmocollin, contributes to cell-cell adhesion. J. Cell Biol. 138, 193-201.

Cuvelier, D., Rossier, O., Bassereau, P. and Nassoy, P. (2003). Micropatterned‘‘adherent/repellent’’ glass surfaces for studying the spreading kinetics ofindividual red blood cells onto protein-decorated substrates. Eur. Biophys. J. 32,342-354.

Delva, E., Tucker, D. K. and Kowalczyk, A. P. (2009). The desmosome. ColdSpring Harb. Perspect. Biol. 1, a002543.

Drees, F., Reilein, A. and Nelson, W. J. (2005). Cell-adhesion assays: fabricationof an E-cadherin substratum and isolation of lateral and Basal membranepatches. Methods Mol. Biol. 294, 303-320.

Dusek, R. L., Godsel, L. M. and Green, K. J. (2007). Discriminating roles ofdesmosomal cadherins: beyond desmosomal adhesion. J. Dermatol. Sci. 45, 7-21.

Eshkind, L., Tian, Q., Schmidt, A., Franke, W. W., Windoffer, R. and Leube,R. E. (2002). Loss of desmoglein 2 suggests essential functions for early

embryonic development and proliferation of embryonal stem cells. Eur. J. CellBiol. 81, 592-598.

Evans, E. and Ritchie, K. (1997). Dynamic strength of molecular adhesion bonds.Biophys. J. 72, 1541-1555.

Franke, W. W. (2009). Discovering the molecular components of intercellularjunction – a historical view. Cold Spring Harb. Perspect. Biol. 1, a003061.

Fuhrmann, A., Getfert, S., Fu, Q., Reimann, P., Lindsay, S. and Ros, R. (2012).Long lifetime of hydrogen-bonded DNA basepairs by force spectroscopy.Biophys. J. 102, 2381-2390.

Gaertner, A., Klauke, B., Stork, I., Niehaus, K., Niemann, G., Gummert, J. andMilting, H. (2012). In vitro functional analyses of arrhythmogenic rightventricular cardiomyopathy-associated desmoglein-2-missense variations.PLoS ONE 7, e47097-e47097.

Garrod, D. R. (1995). Desmosomes and cancer. Cancer Surv. 24, 97-111.Garrod, D. and Kimura, T. E. (2008). Hyper-adhesion: a new concept in cell-celladhesion. Biochem. Soc. Trans. 36, 195-201.

Gehmlich, K., Syrris, P., Peskett, E., Evans, A., Ehler, E., Asimaki, A.,Anastasakis, A., Tsatsopoulou, A., Vouliotis, A.-I., Stefanadis, C. et al.(2011). Mechanistic insights into arrhythmogenic right ventricularcardiomyopathy caused by desmocollin-2 mutations. Cardiovasc. Res. 90, 77-87.

Getsios, S., Amargo, E. V., Dusek, R. L., Ishii, K., Sheu, L., Godsel, L. M. andGreen, K. J. (2004). Coordinated expression of desmoglein 1 and desmocollin 1regulates intercellular adhesion. Differentiation 72, 419-433.

Getsios, S., Simpson, C. L., Kojima, S., Harmon, R., Sheu, L. J., Dusek, R. L.,Cornwell, M. and Green, K. J. (2009). Desmoglein 1-dependent suppression ofEGFR signaling promotes epidermal differentiation and morphogenesis. J. CellBiol. 185, 1243-1258.

Green, K. J. and Simpson, C. L. (2007). Desmosomes: new perspectives on aclassic. J. Invest. Dermatol. 127, 2499-2515.

Green, K. J., Kowalczyk, A. P., Bornslaeger, E. A., Palka, H. L. and Norvell,S. M. (1998). Desmosomes: integrators of mechanical integrity in tissues. Biol.Bull. 194, 374-376, discussion 376-377.

Harmon, R. M., Simpson, C. L., Johnson, J. L., Koetsier, J. L., Dubash, A. D.,Najor, N. A., Sarig, O., Sprecher, E. and Green, K. J. (2013). Desmoglein-1/Erbin interaction suppresses ERK activation to support epidermal differentiation.J. Clin. Invest. 123, 1556-1570.

Harrison, O. J., Bahna, F., Katsamba, P. S., Jin, X., Brasch, J., Vendome, J.,Ahlsen, G., Carroll, K. J., Price, S. R., Honig, B. et al. (2010). Two-stepadhesive binding by classical cadherins. Nat. Struct. Mol. Biol. 17, 348-357.

Hartlieb, E., Kempf, B., Partilla, M., Vigh, B., Spindler, V. and Waschke, J.(2013). Desmoglein 2 is less important than desmoglein 3 for keratinocytecohesion. PLoS ONE 8, e53739.

He, W., Cowin, P. and Stokes, D. L. (2003). Untangling desmosomal knots withelectron tomography. Science 302, 109-113.

Heuser, A., Plovie, E. R., Ellinor, P. T., Grossmann, K. S., Shin, J. T., Wichter,T., Basson, C. T., Lerman, B. B., Sasse-Klaassen, S., Thierfelder, L. et al.(2006). Mutant desmocollin-2 causes arrhythmogenic right ventricularcardiomyopathy. Am. J. Hum. Genet. 79, 1081-1088.

Hutter, J. L. and Bechhoefer, J. (1993). Calibration of atomic-force microscopetips. Rev. Sci. Instrum. 64, 1868.

Kottke, M. D., Delva, E. and Kowalczyk, A. P. (2006). The desmosome: cellscience lessons from human diseases. J. Cell Sci. 119, 797-806.

Kowalczyk, A. P., Borgwardt, J. E. and Green, K. J. (1996). Analysis ofdesmosomal cadherin-adhesive function and stoichiometry of desmosomalcadherin-plakoglobin complexes. J. Invest. Dermatol. 107, 293-300.

Kowalczyk, A. P., Bornslaeger, E. A., Borgwardt, J. E., Palka, H. L., Dhaliwal,A. S., Corcoran, C. M., Denning, M. F. and Green, K. J. (1997). The amino-terminal domain of desmoplakin binds to plakoglobin and clusters desmosomalcadherin-plakoglobin complexes. J. Cell Biol. 139, 773-784.

Marcozzi, C., Burdett, I. D., Buxton, R. S. and Magee, A. I. (1998). Coexpressionof both types of desmosomal cadherin and plakoglobin confers strongintercellular adhesion. J. Cell Sci. 111, 495-509.

Marshall, B. T., Sarangapani, K. K., Wu, J., Lawrence, M. B., McEver, R. P. andZhu, C. (2006). Measuring molecular elasticity by atomic force microscopecantilever fluctuations. Biophys. J. 90, 681-692.

Nekrasova, O. E., Amargo, E. V., Smith, W. O., Chen, J., Kreitzer, G. E. andGreen, K. J. (2011). Desmosomal cadherins utilize distinct kinesins forassembly into desmosomes. J. Cell Biol. 195, 1185-1203.

Nie, Z., Merritt, A., Rouhi-Parkouhi, M., Tabernero, L. and Garrod, D. (2011).Membrane-impermeable cross-linking provides evidence for homophilic,isoform-specific binding of desmosomal cadherins in epithelial cells. J. Biol.Chem. 286, 2143-2154.

Nishizawa, M., Takoh, K. and Matsue, T. (2002). Micropatterning of HeLa Cellson Glass Substrates and Evaluation of Respiratory Activity UsingMicroelectrodes. Langmuir 18, 3645-3649.

North, A. J., Bardsley, W. G., Hyam, J., Bornslaeger, E. A., Cordingley, H. C.,Trinnaman, B., Hatzfeld, M., Green, K. J., Magee, A. I. and Garrod, D. R.(1999). Molecular map of the desmosomal plaque. J. Cell Sci. 112, 4325-4336.

Oda, H. and Takeichi, M. (2011). Evolution: structural and functional diversity ofcadherin at the adherens junction. J. Cell Biol. 193, 1137-1146.

Pasdar, M. and Nelson, W. J. (1988a). Kinetics of desmosome assembly inMadin-Darby canine kidney epithelial cells: temporal and spatial regulation ofdesmoplakin organization and stabilization upon cell-cell contact. I. Biochemicalanalysis. J. Cell Biol. 106, 677-685.

RESEARCH ARTICLE Journal of Cell Science (2014) 127, 2339–2350 doi:10.1242/jcs.146316

2349

Jour

nal o

f Cel

l Sci

ence

Pasdar, M. and Nelson, W. J. (1988b). Kinetics of desmosome assembly inMadin-Darby canine kidney epithelial cells: temporal and spatial regulation ofdesmoplakin organization and stabilization upon cell-cell contact. II.Morphological analysis. J. Cell Biol. 106, 687-695.

Pasdar, M. and Nelson, W. J. (1989). Regulation of desmosome assembly inepithelial cells: kinetics of synthesis, transport, and stabilization of desmoglein I,a major protein of the membrane core domain. J. Cell Biol. 109, 163-177.

Pasdar, M., Krzeminski, K. A. and Nelson, W. J. (1991). Regulation ofdesmosome assembly in MDCK epithelial cells: coordination of membranecore and cytoplasmic plaque domain assembly at the plasma membrane. J. CellBiol. 113, 645-655.

Patel, S. D., Chen, C. P., Bahna, F., Honig, B. and Shapiro, L. (2003). Cadherin-mediated cell-cell adhesion: sticking together as a family. Curr. Opin. Struct.Biol. 13, 690-698.

Pollard, T. D. and Borisy, G. G. (2003). Cellular motility driven by assembly anddisassembly of actin filaments. Cell 112, 453-465.

Rakshit, S., Zhang, Y., Manibog, K., Shafraz, O. and Sivasankar, S. (2012).Ideal, catch, and slip bonds in cadherin adhesion. Proc. Natl. Acad. Sci. USA109, 18815-18820.

Ray, C., Brown, J. R. and Akhremitchev, B. B. (2007). Correction of systematicerrors in single-molecule force spectroscopy with polymeric tethers by atomicforce microscopy. J. Phys. Chem. B 111, 1963-1974.

Roberts, G. A., Burdett, I. D., Pidsley, S. C., King, I. A., Magee, A. I. andBuxton, R. S. (1998). Antisense expression of a desmocollin gene in MDCKcells alters desmosome plaque assembly but does not affect desmogleinexpression. Eur. J. Cell Biol. 76, 192-203.

Runswick, S. K., O’Hare, M. J., Jones, L., Streuli, C. H. and Garrod, D. R.(2001). Desmosomal adhesion regulates epithelial morphogenesis and cellpositioning. Nat. Cell Biol. 3, 823-830.

Saffitz, J. E., Jongbloed, J. D. H., van der Smagt, J., Lombardi, P. M.,Wiesfeld, A. C. P., Nelen, M., Schouten, M., Jongbloed, R., Cox, M. G. P. J.,van Wolferen, M. et al. (2009). Desmosome mutations in arrhythmogenic rightventricular cardiomyopathy: important insight but only part of the picture. Circ.Cardiovasc. Genet. 2, 415-417.

Saito, M., Tucker, D. K., Kohlhorst, D., Niessen, C. M. and Kowalczyk, A. P.(2012). Classical and desmosomal cadherins at a glance. J. Cell Sci. 125, 2547-2552.

Schlegel, N., Meir, M., Heupel, W.-M., Holthofer, B., Leube, R. E. and Waschke,J. (2010). Desmoglein 2-mediated adhesion is required for intestinal epithelialbarrier integrity. Am. J. Physiol. 298, G774-G783.

Scott, D. W. (1979). On optimal and data-based histograms. Biometrika 66, 605-610.

Shapiro, L. and Weis, W. I. (2009). Structure and biochemistry of cadherins andcatenins. Cold Spring Harb. Perspect. Biol. 1, a003053-a003053.

Shapiro, L., Fannon, A. M., Kwong, P. D., Thompson, A., Lehmann, M. S.,Grubel, G., Legrand, J. F., Als-Nielsen, J., Colman, D. R. and Hendrickson,W. A. (1995). Structural basis of cell-cell adhesion by cadherins. Nature 374,327-337.

Simpson, M. A., Mansour, S., Ahnood, D., Kalidas, K., Patton, M. A.,McKenna, W. J., Behr, E. R. and Crosby, A. H. (2009). Homozygousmutation of desmocollin-2 in arrhythmogenic right ventricular cardiomyopathywith mild palmoplantar keratoderma and woolly hair. Cardiology 113, 28-34.

Simpson, C. L., Patel, D. M. and Green, K. J. (2011). Deconstructing the skin:cytoarchitectural determinants of epidermal morphogenesis. Nat. Rev. Mol. CellBiol. 12, 565-580.

Sivasankar, S., Zhang, Y., Nelson, W. J. and Chu, S. (2009). Characterizing theinitial encounter complex in cadherin adhesion. Structure 17, 1075-1081.

Smith, S. B., Finzi, L. and Bustamante, C. (1992). Direct mechanicalmeasurements of the elasticity of single DNA molecules by using magneticbeads. Science 258, 1122-1126.

Spiliotis, E. T., Kinoshita, M. and Nelson, W. J. (2005). A mitotic septin scaffoldrequired for Mammalian chromosome congression and segregation. Science307, 1781-1785.

Syed, S.-E.-H., Trinnaman, B., Martin, S., Major, S., Hutchinson, J. and Magee,A. I. (2002). Molecular interactions between desmosomal cadherins. Biochem.J. 362, 317-327.

Thomason, H. A., Scothern, A., McHarg, S. and Garrod, D. R. (2010).Desmosomes: adhesive strength and signalling in health and disease. Biochem.J. 429, 419-433.

Troyanovsky, S. M., Troyanovsky, R. B., Eshkind, L. G., Krutovskikh, V. A.,Leube, R. E. and Franke, W. W. (1994a). Identification of the plakoglobin-binding domain in desmoglein and its role in plaque assembly and intermediatefilament anchorage. J. Cell Biol. 127, 151-160.

Troyanovsky, S. M., Troyanovsky, R. B., Eshkind, L. G., Leube, R. E. andFranke, W. W. (1994b). Identification of amino acid sequence motifs indesmocollin, a desmosomal glycoprotein, that are required for plakoglobinbinding and plaque formation. Proc. Natl. Acad. Sci. USA 91, 10790-10794.

Waschke, J. (2008). The desmosome and pemphigus. Histochem. Cell Biol. 130,21-54.

Yamada, S., Pokutta, S., Drees, F., Weis, W. I. and Nelson, W. J. (2005).Deconstructing the cadherin-catenin-actin complex. Cell 123, 889-901.

Zhang, Y., Sivasankar, S., Nelson, W. J. and Chu, S. (2009). Resolving cadherininteractions and binding cooperativity at the single-molecule level. Proc. Natl.Acad. Sci. USA 106, 109-114.

RESEARCH ARTICLE Journal of Cell Science (2014) 127, 2339–2350 doi:10.1242/jcs.146316

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