David B. Williams

David B. Williams


BSc, University of Manitoba, 1973
MSc, University of Toronto, 1975
PhD, University of Toronto, 1981
Postdoc, Johns Hopkins University, 1981-1984

Address Medical Sciences Building, Room 5316
Toronto, ON M5S 1A8
Lab Phone 416-978-6034
Office Phone 416-978-2546
Email david.williams@utoronto.ca

David Williams did his doctoral work with Harry Schachter at the University of Toronto in the area of glycoprotein biogenesis followed by postdoctoral studies at Johns Hopkins University with Bill Lennarz and Gerald Hart where he developed an interest in protein folding and quality control in the mammalian secretory pathway. As a principal investigator he discovered the calnexin chaperone system and has characterized it extensively. He has also made important contributions to the field of antigen presentation by class I histocompatibility molecules, particularly in how these molecules assemble with antigenic peptides in the endoplasmic reticulum. Current research focuses on the roles of molecular chaperones, protein disulfide isomerases and peptidyl prolyl isomerases in protein folding as well as in the quality control triage system of the endoplasmic reticulum.

He has held several administrative positions within the Biochemistry Department including Acting Chair (1996-1997) and Graduate Coordinator (2001-2005). He was also elected President of the Canadian Society of Biochemistry, Molecular and Cellular Biology in 2009 and has served on the Medical Review Panel of the Gairdner Award Foundation from 2009 – 2013.

In the News

Research Lab

The Williams Lab is based within the Medical Sciences Building in proximity to a number of laboratories studying protein folding as well as the mechanisms of action of molecular chaperones. As such, it provides an excellent environment for interactions between trainees working in related fields. We use both in vitro and cell biological approaches to study the molecules involved in protein folding and quality control processes within the endoplasmic reticulum. Trainees are exposed to a variety of techniques and approaches including protein purification, site-directred mutagenesis, in vitro aggregation and folding assays, cell culture, transfection, protein depletion by RNA interference, protein immunoisolation/immunoblotting, and pulse-chase cell radiolabeling.


We have ongoing collaborations with a number of excellent research groups including:

Dr. Walid Houry, University of Toronto
Dr. Kalle Gehring, McGill University (protein structure by X-ray crystallography and NMR spectroscopy)
Dr. David Westaway, University of Alberta (cell and animal models of prion diseases)

Williams Lab 2014

Williams Lab 2014

Current Lab Members

Daniel Chapman, PhD student
Max Sawicki, MSc student
Debbie Hong, MSc Student
Samar Ahmad, MSc student (visiting from University of Oulu, Finland)

Dr. Williams is not currently accepting new graduate students or postdoctoral fellows.

Research Description

Protein Folding & Quality Control within the Endoplasmic Reticulum

Our research focuses on how proteins fold and assemble within the endoplasmic reticulum (ER), including the cellular machinery that monitors the integrity of this process. Protein folding within the ER is complex and highly coordinated, involving post-translational modifications such as Asn-linked glycosylation and disulfide bond formation as well as the acquisition of native tertiary and quaternary structure. Furthermore, the ER houses a sophisticated “quality control” system that monitors the folding status of a protein and retains misfolded conformers for eventual destruction by a process known as ER-associated degradation (ERAD). Many inherited human diseases such as cystic fibrosis involve ER retention and disposal of aberrant proteins. Since these proteins often retain a degree of function, there is intense interest in studying ER folding and quality control to learn how these processes may be manipulated in disease states to enhance trafficking of mutant proteins to their normal sites of action and improve function. Our efforts are directed toward characterizing major players in ER folding and quality control. These include several molecular chaperones, a variety of folding enzymes, as well as components of ERAD disposal pathways.

Molecular Chaperones

Molecular chaperones bind transiently to hydrophobic segments of nascent polypeptide chains thereby preventing aggregation and allowing productive folding to occur more efficiently. Our focus is on two molecular chaperones of the ER, membrane-bound calnexin and its soluble paralogue, calreticulin. Calnexin and calreticulin recognize newly synthesized proteins in a remarkable way. Both chaperones possess a lectin site that allows them to bind to Asn-linked oligosaccharides that have a single terminal glucose residue, as well as a polypeptide binding site that recognizes non-native polypeptide chains. Furthermore, calnexin and calreticulin bind to two folding catalysts within the ER, the protein disulfide isomerase family member ERp57 and the cis-trans prolyl isomerase cyclophilin B (CypB). These complexes are thought to extend the chaperone functions of calnexin/calreticulin by promoting disulfide bond formation or catalyzing proline isomerization in the glycoproteins that bind to these chaperones.


Current work is directed toward determining the relative contributions of the lectin, polypeptide, and ERp57/CypB binding sites to the overall functions of these chaperones. We do this by site-directed mutagenesis of each site and assessing the consequences in models of protein folding both in vitro, using entirely purified components, and by expressing the mutants in cultured cells. We are also interested in whether calnexin, calreticulin and other ER chaperones retain mutant proteins associated with human diseases. Interfering with chaperone interactions may enhance export of these mutants from the ER

Protein disulfide isomerases (PDIs)

The formation of disulfide bonds in newly synthesized proteins, the reshuffling of incorrect disulfides, and the removal of disulfides during ERAD disposal of misfolded proteins are reactions catalyzed by the protein disulfide isomerases. Remarkably, there are more than 20 PDI family members in the human ER and a major question is …. why so many? Yeast manage very well with just 5! We are approaching this issue by systematically depleting PDIs, individually and in combination, using high-efficiency RNA interference. We then monitor the consequences on disulfide formation in newly synthesized proteins as well as on the reduction of disulfides during ERAD disposal of misfolded proteins. We are discovering that, rather than simply possessing redundant activities, there is a division of labour among family members; some catalyze disulfide formation, others remove disulfides during ERAD disposal, and others have distinct substrate specificity characteristics.

Peptidyl prolyl isomerases (PPIs)

These interesting enzymes catalyze the interconversion of cis and trans isomers of the peptide bond preceding proline residues. Within the cytosol, interconversion of cis– and trans-Pro isomers typically functions as a molecular switch, controlling two protein conformers that may have distinct functions or bind to distinct partners. Within the ER, there are at least 5 PPIs, called cyclophilin B (CypB) and FKBP13, 23, 63 and 65, but their functions are poorly understood. They may catalyze the rate-limiting formation of cis-Pro bonds in nascent proteins, regulate the activities of cis-Pro-containing proteins (as in the cytosol), or remove the tight bends associated with cis-Pro bonds during ERAD disposal of misfolded proteins. We are taking the same systematic knockdown approach that we are using for the PDIs and are discovering important roles for the peptidyl prolyl isomerases both in protein folding and in unfolding/degradation pathways.

ER-associated degradation (ERAD)

ERADAs part of its quality control system, the ER disposes of misfolded proteins in an unusual manner. Such proteins are targeted to a retrotranslocation complex where they are unfolded and translocated across the ER membrane into the cytosol for degradaton by the multi-catalytic proteasome complex. It has become apparent that multiple ERAD pathways exist, in part based on the topology of the misfolded protein (soluble, membrane-bound, polytopic, etc.). These pathways are incompletely characterized. To gain insights into their composition and mechanism of action, we have undertaken a genome-wide shRNA knockdown screen to identify novel ERAD components. This has been remarkably successful and we are now characterizing the many intriguing “hits” obtained. Not surprisingly, these include members of the chaperone, PDI, and PPI families as well as components of the ubiquitin-proteasome degradation machinery.

Awards & Distinctions

1994 — Merck Frost Prize for outstanding initial independent research in Biochemistry and Molecular Biology in Canada
2000 — University of Toronto Dales Award for sustained excellence in medical research
2009 — President of the Canadian Society of Biochemistry, Molecular and Cellular Biology

Courses Taught

BCH478H Advanced Biochemistry Lab
BCH444H Protein Trafficking in the Secretory and Endocytic Pathways

Extra-Departmental Courses

IMM1016H Recent Advances in Basic Immunology


View all publications on PubMed

Contributions of the lectin and polypeptide binding sites of calreticulin to Its chaperone functions in vitro and in cells.
Lum, R., Ahmad, S., Hong, S-J., Chapman, D.C., Kozlov, G. and Williams, D.B.
J. Biol. Chem. 2016. 291(37):19631-41.  Read

Inhibition of the FKBP family of peptidyl prolyl isomerases induces abortive translocation and degradation of the cellular prion protein.
Stocki, P., Sawicki, M., Mays, C. E., Hong, S. J., Chapman, D. C., Westaway, D. and Williams, D. B.
Mol. Biol. Cell 2016. 27(5):757-767.  Read

Cyclophilin C participates in the US2-mediated degradation of major histocompatibility complex Class I molecules.
Chapman, D. C., Stocki, P. and Williams, D.B.
PLoS One. 2015. 10(12):e0145458.  Read

Depletion of cyclophilins B and C leads to dysregulation of endoplasmic reticulum redox homeostasis.
Stocki P, Chapman DC, Beach LA, Williams DB
J Biol Chem. 2014. 289(33):23086-96  Read

Vitamin K epoxide reductase contributes to protein disulfide formation and redox homeostasis within the endoplasmic reticulum.
Rutkevich LA, Williams DB.
Mol Biol Cell. 2012. 23(11):2017-27  Read

Structural and functional relationships between the lectin and arm domains of calreticulin.
Pocanschi, C.L., Kozlov, G., Brockmeier, U., Brockmeier, A., Williams, D. B. and Gehring, K.
J. Biol. Chem. 2011. 286: 27266-27277  Read

Participation of lectin chaperones and thiol oxidoreductases in protein folding within the endoplasmic reticulum.
Rutkevich, L.A. and Williams, D.B.
Curr. Opin. Cell Biol. 2011. 23(2):157-166.  Read

Functional relationship between protein disulfide isomerase family members during the oxidative folding of human secretory proteins.
Rutkevich, L. A., Cohen-Doyle, M. F., Brockmeier, U. and Williams, D. B.
Mol. Biol. Cell 2010. 21: 3093-3105.  Read

Structural basis of carbohydrate recognition by calreticulin.
Kozlov G, Pocanschi CL, Rosenauer A, Bastos-Aristizabal S, Gorelik A, Williams DB, Gehring K.
J. Biol. Chem. 2010. 285:38612-38620  Read