Naa-Adjeley Ablorh

Graduate student (Biochemistry)

Office:	1-152 Nils Hasselmo Hall
Phone:	(612) 626-0113
Fax:	(612) 624-0632
Mail:	Dept. of Biochemistry (BMBB) U of Minnesota 6-155 Jackson Hall 321 Church St. Minneapolis, MN 55455
Email:	naa@ddt.biochem.umn.edu

Research Interests:
PLB phosphorylation

Sarah Blakely Anderson

Assistant Scientist - Lab Manager

Office:	5-101C Nils Hasselmo Hall
Phone:	(612) 624-9918
Fax:	(612) 624-0632
Mail:	Dept. of Biochemistry (BMBB) University of Minnesota 6-155 Jackson Hall 321 Church St. Minneapolis, MN 55455
Email:	sb@ddt.biochem.umn.edu

Administrative

Manage lab of approximately 35 employees (professors, research assistant professors, post-doctoral associates, graduate students, technical staff, and undergraduate students). Manage spending and salary distribution on 26 grants, approximately $1.5M annual budget. Place supply and equipment orders, justify all purchases, and properly charge grants. Act as liaison between office administrative (accountants and HR) and lab staff. Work with Office of Human Resources and Medical School to get hiring approval, write and position postings for new hires, interview and advise on technical staff hires. Hire, supervise, and train undergraduate student workers. Orient new employees to lab procedures, document safety training, and request UCard access to building. Schedule weekly lab meetings, keep lab calendar current. Organize lab help document (DDTLABHELP), maintain hyperlinks, keep protocols current, and recruit other responsible lab members to document their protocols.

Wetlab

Monitor lab inventory and order all supplies and equipment. Maintain service contracts on all equipment, schedule service calls, and coordinate repairs. Write and maintain research compliance protocols (IACUC, IRB, and IBC). Oversee semi-annual animal and hazardous waste inspections with various regulatory offices. Serve as lab safety and controlled substance officer. Provide annual safety and chemical training to all employees, and oversee all waste compliance in lab. Act as point-of-contact for any safety or chemical related problems and communicates with Dept of Environmental Health and Safety to resolve problems.

Scientific

Assess protein needs for muscle group. Coordinate and organize protein preps and document in mutant database. Advise lab members in molecular biology. Aid in the creation of new myosin mutants.

Research Techniques

Molecular biology (restriction digest, ligation, plasmid purification)
Site-directed mutagenesis
PCR
Cloning / subcloning
Cell culture (E.coli, Spodoptera frugiperda, Dictyostelium discoideum)
Sterile technique
Protein expression (E.coli, Spodoptera frugiperda, Dictyostelium discoideum)
Western blots
Gel electrophoresis (SDS-PAGE, agarose, urea)
Gel purification (DNA)
Protein purification (native and expressed)
Column Chromatography (anion exchange, size exclusion, affinity)
Spectroscopic labeling (Electron Paramagnetic Resonance, Fluorescence)
UV/Vis spectrophotometry
Protein concentration assays (Bradford, BCA)
ATPase activity assays (Malachite Green method)
Fiber dissection (rabbit, scallop)

Publications

Nesmelov, Y.E., R. Agafonov, I.V. Negrashov, S. Blakely, M.A. Titus, and D.D. Thomas. 2011. Structural kinetics of myosin by transient time-resolved FRET. Proc Nat Acad Sci USA 108:1891�1896.

Agafonov R, I. V. Negrashov, Y. Tkachev, S. Blakely, M. A. Titus, D. D. Thomas, and Y. E. Nesmelov. 2009. Structural dynamics of the myosin relay helix by time-resolved EPR and FRET. Proc Nat Acad Sci USA. 106: 21625-21630.

Korman, V.L., S.E. Anderson, E. Prochniewicz, M.A. Titus, and D.D. Thomas. 2006. Structural dynamics of the actin-myosin interface by site-directed spectroscopy. J Mol Biol. 356: 1107-1117.

Nelson, W.D., S.E. Blakely, Y.E. Nesmelov, and D.D. Thomas. 2005. Site-directed spin labeling reveals a conformational switch in the phosphorylation domain of smooth muscle myosin. Proc Natl Acad Sci U S A. 102: 4000-4005. PMCID: 554790.

Chris Arcand

Network Administrator

Office:	1-138 Nils Hasselmo Hall
Phone:	(612) 625-9196
Fax:	(612) 624-0632
Mail:	Dept. of Biochemistry (BMBB) University of Minnesota 6-155 Jackson Hall 321 Church St. Minneapolis, MN 55455
Email:	cpa@ddt.biochem.umn.edu

Mike Autry

Research Associate

Office:	1-138 Nils Hasselmo Hall
Phone:	(612) 626-3225
Fax:	(612) 624-0632
Mail:	Dept. of Biochemistry (BMBB) University of Minnesota 6-155 Jackson Hall 321 Church St. Minneapolis, MN 55455
Email:	jma@ddt.biochem.umn.edu

Research Interests

I am investigating the structural dynamics of the sarcoplasmic reticulum Ca2+-transporting ATPase (SERCA), which is responsible for mediating muscle relaxation by removing calcium from the actin/myosin molecular motors inside the muscle cell. I am also investigating the functional interaction of SERCA with sarcolipin and phospholamban, two small integral membrane proteins that regulate calcium transport and muscle contractility in a tissue-specific manner. Techniques: cDNA cloning, protein mutagenesis, baculovirus-insect cell expression, recombinant protein purification, functional reconstitution, site-directed fluorescence spectroscopy.

Publications

Peer-reviewed:

1. Simmerman HKB, Kobayashi YM, Autry JM, Jones LR. A leucine zipper stabilizes the pentameric membrane domain of phospholamban and forms a coiled-coil pore structure. The Journal of Biological Chemistry 271:5941-5946, 1996� [136 citations]

2. Cornea RL, Jones LR, Autry JM, Thomas DD. Mutation and phosphorylation change the oligomeric structure of phospholamban in lipid bilayers. Biochemistry 36:2960-2967, 1997 [82 citations]

3. Autry JM, Jones LR. Functional co-expression of the canine cardiac Ca2+ pump and � phospholamban in Spodoptera frugiperda (Sf21) cells reveals new insights on ATPase regulation. The Journal of Biological Chemistry 272:15872-15880, 1997� [73 citations]

4. Li M, Cornea RL, Autry JM, Jones LR, Thomas DD. Phosphorylation-induced structural change in phospholamban and its mutants, detected by intrinsic fluorescence. Biochemistry 37:7869-7877, 1998� [16 citations]

5. Reddy LG, Autry JM, Jones LR, Thomas DD. Co-reconstitution of phospholamban mutants with the Ca-ATPase reveals dependence of inhibitory function on phospholamban structure. The Journal of Biological Chemistry 274:7649-7655, 1999 [29 citations]

6. Karon BS, Autry JM, Shi Y, Garnett CE, Inesi G, Jones LR, Kutchai H, Thomas DD. Different anesthetic sensitivities of skeletal and cardiac isoforms of the Ca-ATPase. Biochemistry 37:9301-9307, 1999� [12 citations]

7. Cornea RL, Autry JM, Chen Z, Jones LR. Re-examination of the role of the leucine/isoleucine zipper residues of phospholamban in inhibition of the calcium pump of cardiac sarcoplasmic reticulum. The Journal of Biological Chemistry 275:41487-41494, 2000� [22 citations]

8. Mahaney JE, Autry JM, Jones LR. Kinetic studies of the cardiac Ca-ATPase expressed in Sf21 cells: New insights on Ca-ATPase regulation by phospholamban. Biophysical Journal 78:1306-1323, 2000� [5 citations]

Conference Proceedings:

1. Autry JM, Jones LR. High-level coexpression of the canine cardiac calcium pump and phospholamban in Sf21 insect cells. Annals of the New York Academy of Sciences. 853:92-102, 1998 [7 citations]

2. Thomas DD, Reddy LG, Karim CB, Li M, Cornea R, Autry JM, Jones LR, Stamm J. Direct spectroscopic detection of molecular dynamics and interactions of the calcium pump and phospholamban. Annals of the New York Academy of Sciences. 853:186-194, 1998� [22 citations]

Other Articles:

1. Klotz DM, Autry JM, Clark RJ. NPA Policy Committee Monthly Report, 2005

2. Autry JM. NRC “Bridges to Independence” Report. UMN-PDA News, Spring 2005

3. Autry JM. NRC Report on NIH Research Training Programs. UMN-PDA News, Summer 2005

4. Autry JM. Recommended Departmental Practices: Giving Postdocs Order. NPA POSTDOCket, Summer 2005

5. Featured interview in Science Next Wave: “Making it Great for Everybody” by Beryl Benderly, Summer 2005

GenBank Sequences:

1. Autry JM, Scott BT, Jones LR. U94345: Cloning of the cardiac Ca2+-ATPase cDNA (SERCA2a) from dog left ventricle using a ?gt10 cDNA library (1997)

2. Autry JM, Thomas DD. Cloning of the fast-twitch Ca2+-ATPase cDNA (SERCA1a) from rabbit psoas muscle using RT-PCR (waiting for publication)

3. Autry JM, Thomas DD. Cloning of the slow-twitch Ca2+-ATPase cDNA (SERCA2a) from rabbit soleus muscle using RT-PCR (waiting for publication)

4. Autry JM, Thomas DD. Cloning of the cardiac Ca2+-ATPase cDNA (SERCA2a) from dog left ventricle using RT-PCR (waiting for publication)

5. Autry JM, Thomas DD. Cloning of phospholamban cDNA (PLB) from dog left ventricle using RT-PCR (waiting for publication)

6. Autry JM, Thomas DD. Cloning of sarcolipin cDNA (SLN) from rabbit psoas muscle using RT-PCR (waiting for publication)

Acknowledgements:

Christine Karim, Biochemistry 1998

Zhenhui Chen, Doctoral Thesis, Indiana University 1999

Jason Waggoner, Protein Expression and Purification 2003

Seth Robia, Biochemistry 2005

Recent Poster Presentations:

1. Kirchhefer U (presenter), Autry JM, Jones LR, Schmitz W, Neumann J. Cardiac-specific overexpression of the native and a double mutant form of SERCA2a in transgenic mice results in different biochemical and physiological phenotypes. Deutsche Gesellschaft f�r Experimentelle und Klinische, Mainz Germany (2004)

2. Autry JM, Winters DL, Juhl ME, Thomas DD. Overproduction of SERCA tetra-cysteine mutants for site-directed labeling with biarsenical fluorophores. Biophysical Society Meeting, Long Beach CA (2005)

3. Autry JM, Brennan RE, VanDrasek B, Sivasubbu S, Walsh R, Balciuniene J, Cheong A, El-Fakahany EE. The University of Minnesota Postdoctoral Association: A helping hand for enhancing the postdoctoral experience. National Postdoctoral Association Meeting, San Diego CA (2005)

4. Winters DL (presenter), Autry JM, Thomas DD. Functional dynamics of SERCA as revealed by cyan fluorescent protein (CFP) fused to the actuator domain. 11th International Na,K-ATPase Conference, Woods Hole MA (Sept 2005)

5. Autry JM*, Winters DL, Thomas DD. New methods for site-directed spectroscopic labeling of SERCA-type Ca-ATPases. 11th International Na,K-ATPase Conference, Woods Hole MA (Sept 2005) *received Symposium Travel Award

Manuscripts in Preparation (Autry as first author):

1. Autry JM, Thomas DD, et al. Targeted mutagenesis of the phospholamban transmembrane domain identifies novel structural determinants in pentamer formation and functional regulation of the cardiac calcium pump (6100 words, 12 figures)

2. Autry JM, Thomas DD, et al. Baculovirus expression and affinity purification of recombinant SERCA1 with high specific activity (3000 words, 4 �figs)

3. Autry JM, Thomas DD, et al. Structural dynamics of SERCA using bis-arsenical fluorescein (FLASH) rigidly coupled to an engineered tetra-cysteine motif in the phosphorylation domain (2600 words, 7 figures)

4. Autry JM, Thomas DD, et al. In vivo oligomerization of phospholamban and sarcolipin (4400 words; 9 figures)

Ben Binder

BMBB Rotation Student

Office:	5-290 Nils Hasselmo Hall
Phone:	(612) 626-3322
Fax:	(612) 624-0632
Mail:	Dept. of Biochemistry (BMBB) University of Minnesota 6-155 Jackson Hall 321 Church St. Minneapolis, MN 55455
Email:	bpb@ddt.biochem.umn.edu

Research Interests

The generation of force in muscle contraction, while complex and nuanced in its full consideration, is nonetheless the direct result of a single interaction between two proteins, actin and myosin. This interaction represents the crucial device responsible for the shortening of muscle fibers, and in that partnership myosin takes the active role, unlocking the energy stored in ATP to pull on actin filaments. While a broad understanding of this force-generating cycle has been established for many years (Fig. 1), recent evidence suggests that myosin does not follow a progression of discreet structural transitions throughout the binding and hydrolysis of ATP, but rather constantly entertains a delicate balance of structural states within a population, achieving its function through subtle shifts in conformational equilibria.

The derivative of ATP bound to myosin at a given step in the hydrolytic cycle (myosin's "biochemical state") has long been associated with a particular protein conformation (myosin's "structural state"), but our lab has shown through spectroscopic analysis that populations with homogeneous biochemical states consistently display marked structural heterogeneity. This finding has important implications not only for the challenge of describing myosin's normal behavior, but also for understanding the fundamental nature of various disease phenotypes. My research focuses on providing new insight into the structural dynamics of myosin, using spectroscopy to characterize how distributions of structural states are affected by different physiological conditions.

Spectroscopic Investigation of Oxidative Stress in Myosin

Reactive oxygen species (ROS) are produced as by-products of cellular respiration, and normally exist as essential components in a carefully-controlled redox environment. However, if the balance is preturbed and the proliferation of ROS overwhelms cellular antioxidant systems, the resulting oxidative conditions can lead to unchecked structural modification of proteins. We hypothesize that oxidation of myosin at various functionally-relevant sites contributes to the phenotypes observed in heart failure, muscle degeneration and biological aging, where elevated levels of ROS have also been observed (Fig. 2). Our lab has correlated ROS exposure with a decrease in myosin's ability to generate force both in vitro and in vivo. I seek to expand upon these observations by identifying specific residues on the myosin catalytic domain that impair function when oxidized, and using spectroscopy to observe the effects of site-specific oxidation on structural dynamics.

Ashley Brate

Undergraduate Research Assistant

Research Interests:

My current research interest is looking at the effects of disease-causing point mutations of dystrophin proteins on actin�s resilience.� We have already begun testing full length dystrophin mutants that are described in Henderson, et al.� Under my current project, I am working on making shorter constructs of both dystrophin and utrophin proteins with mutations, expressing them in E. coli cells, and then using these constructs to study the ways in which these proteins are similar or different from each other when they bind to actin.� The information obtained will help aid in muscular dystrophy gene therapy work.

Roy Collins

Senior Network Administrator

Office:	1-138 Nils Hasselmo Hall
Phone:	(612) 625-9196
Fax:	(612) 624-0632
Mail:	Dept. of Biochemistry (BMBB) University of Minnesota 6-155 Jackson Hall 321 Church St. Minneapolis, MN 55455
Email:	rtc@ddt.biochem.umn.edu

Brett Colson

Postdoctoral Fellow

Office:	1-100 Nils Hasselmo Hall
Phone:	(612) 625-6702
Fax:	(612) 624-0632
Mail:	Dept. of Biochemistry (BMBB) University of Minnesota 6-155 Jackson Hall 321 Church St. Minneapolis, MN 55455
Email:	bc@ddt.biochem.umn.edu

CURRICULUM VITAE

Research Interests

I am a structural biophysicist interested in the mechanics of muscle force generation and muscle protein spectroscopy as it relates to molecular, sub-cellular, and organismal physiology. A majority of my research focuses on understanding the relationships between transitions in protein structure and transitions in the force development and relaxation of cardiac muscle. My overreaching goal is to target and therapeutically mimic post-translationally modified states of muscle proteins, in efforts to enhance regulatory and modulatory capabilites of myofilaments in the failing heart, as well as dysfunction in skeletal and smooth muscle systems.

My work focuses on understanding the mechanism by which phosphorylation of muscle regulatory proteins mediate myosin and actin structural dynamics to influence contraction. Specifically, activation of the myosin-actin interaction in smooth muscle by phosphorylation of the regulatory light chain (RLC) and acceleration of the myosin-actin interaction kinetics in cardiac muscle by phosphorylation of myosin binding protein-C (MyBP-C).

Site-directed spectroscopy is a powerful biophysical approach for deciphering molecular motions and interactions of proteins and protein complexes with high resolution in both space and time. After attaching site-directed probes to select muscle proteins, I use time-resolved fluorescence resonance energy transfer (TR-FRET) and time-resolved phosphorescence anisotropy (TPA) to directly detect structure and dynamics of muscle proteins and complexes in solution. I intend to spectroscopically study molecular motions associated with force generation, by using electron paramagnetic resonance (EPR) in muscle fibers, and by using pulsed double electron electron resonance (DEER) in macromolecular complexes especially relevant to cardiovascular diseases.

Publications

Colson, B.A., M.R. Locher, T. Bekyarova, J.R. Patel, D.P. Fitzsimons, T.C. Irving and R.L. Moss (2010). Differential roles of regulatory light chain and myosin binding protein-C phosphorylations in the modulation of cardiac force development. J Physiol 588:981-93.

Colson, B.A., T. Bekyarova, M.R. Locher, D.P. Fitzsimons, T.C. Irving and R.L. Moss (2008). Protein kinase A-mediated phosphorylation of cMyBP-C increases proximity of myosin heads to actin in resting myocardium. Circ Res 103:244-251.

Colson, B.A., T. Bekyarova, D.P. Fitzsimons, T.C. Irving and R.L. Moss (2007).�Radial displacement of myosin cross-bridges in mouse myocardium due to ablation of myosin binding protein-C. J Molec Biol 367:36-41.

Iustin Cornea

Web Administrator

Phone:	(651) 331-6778
Email:	ic@ddt.biochem.umn.edu

Research Interests

My research involves computational modeling of actin-bound conformations of dystrophin. In order to elucidate the open structure of dystrophin when it is bound to actin, our lab has conducted DEER experiments to determine the distances between two pairs of probes. Using these distance contraints and VMD, I have developed a computational method to take a model of dystrophin and manipulate its domains to conform to the aforementioned distance constraints.

With only two constraints, there is still some ambiguity in the exact orientations of the two domains. My method also serves to generate an arbitrary number of analogous models that still conform to the distance constraints, but with varied domain orientations.

Octavian Cornea

IT Professional - Lab Administrative Manager

Office:	5-101A Nils Hasselmo Hall
Phone:	(612) 624-5688
Fax:	(612) 624-0632
Mail:	Dept. of Biochemistry (BMBB) University of Minnesota 6-155 Jackson Hall 321 Church St. Minneapolis, MN 55455
Email:	oc@ddt.biochem.umn.edu

Sinziana Cornea

Junior Scientist

Office:	5-290 Nils Hasselmo Hall
Phone:	(612) 626-3322
Fax:	(612) 624-0632
Mail:	Dept. of Biochemistry (BMBB) University of Minnesota 6-155 Jackson Hall 321 Church St. Minneapolis, MN 55455
Email:	sc@ddt.biochem.umn.edu

Jonathan Crain

Graduate Student (BMBB)

Office:	1-152 Nils Hasselmo Hall
Phone:	(612) 626-0113
Fax:	(612) 624-0632
Mail:	Dept. of Biochemistry (BMBB) University of Minnesota 6-155 Jackson Hall 321 Church St. Minneapolis, MN 55455
Email:	jc@ddt.biochem.umn.edu

Daphne Dong

Graduate Student (BME)

Office:	1-152 Nils Hasselmo Hall
Phone:	(612) 626-0113
Fax:	(612) 624-0632
Mail:	Dept. of Biochemistry (BMBB) University of Minnesota 6-155 Jackson Hall 321 Church St. Minneapolis, MN 55455
Email:	xd@ddt.biochem.umn.edu

Research Interests

My research is focused on the structural basis of the regulation of the sarcoplasmic reticulum Ca-ATPase (SERCA) by its endogenous cardiac inhibitor phospholamban (PLB). SERCA actively transports Ca2+ back into the sarcoplasmic reticulum to induce muscle relaxation. In the failing heart, SERCA's inhibition by PLB increases, resulting in inadequate Ca2+ reuptake and hence deficient muscle relaxation. One of the promising strategies to cure heart failure (HF) is by reducing SERCA inhibition by PLB. My goal is to map the structural and dynamic changes within the SERCA-PLB complex that are needed to relieve inhibition. My principal biophysical method is fluorescence resonance energy transfer (FRET), which measures interprobe distances. By performing time-resolved FRET with a pulsed laser, I can measure distances distributions and resolve complex structural equilibria. My goal is to provide the blueprint needed for the rational design of drug and gene therapies.

L. Michel Espinoza-Fonseca

Postdoctoral Associate

Office:	1-136 Nils Hasselmo Hall
Phone:	(612) 626-0113
Fax:	(612) 624-0632
Mail:	Dept. of Biochemistry (BMBB) University of Minnesota 6-155 Jackson Hall 321 Church St. Minneapolis, MN 55455
Email:	mef@ddt.biochem.umn.edu

Research Interests

Disorders of the cardiac sarcoplasmic reticulum calcium ATPase (SERCA) pump and its regulation by the small transmembrane protein phospholamban (PLB) are central to heart failure, the leading cause of death in the USA, therefore structure-based therapies are  becoming increasingly essential in the effective treatment of heart failure. The focus of my research is to investigate the regulation of SERCA by PLB. I use a combination of computational docking and molecular dynamics (MD) simulations to determine the structure of the SERCA-PLB complex along the catalytic cycle of SERCA. My research interests go beyond creating  flexible models of SERCA-PLB complexes: I am interested in understanding, in atomic detail, the dynamic binding interactions between these membrane proteins. How do these proteins interact in different conformations of SERCA?  How do these interactions affect SERCA dynamics? To answer these questions, I use a combination of state-of-the-art computer simulations and analysis techniques.  I am also interested in the molecular motions of SERCA and the coupling of intrinsic conformational disorder and dynamics as an effective mechanism for regulation of calcium transport in the heart. This work is essential to visualize the implications of experiments at the molecular level and for the rational design of drugs or gene therapy approaches to combat heart failure.

Selected Publications

Espinoza-Fonseca LM*, Ilizaliturri-Flores I, Correa-Basurto J. Backbone conformational preferences of an intrinsically disordered protein in solution. Mol. Biosyst., 2012, 8, 1798�1805.

Espinoza-Fonseca LM*. Dynamic optimization of signal transduction via intrinsic disorder. Mol. Biosyst., 2012,8, 194�197.

Espinoza-Fonseca LM*. Aromatic residues link binding and function of intrinsically disordered proteins. Mol. Biosyst., 2012. 8, 237�246.

Espinoza-Fonseca LM, Thomas DD. Atomic-level characterization of the activation mechanism of SERCA by calcium. PloS ONE, 2011, 6(10): e26936.

Kast D, Espinoza-Fonseca LM, Yi C, Thomas DD. Phosphorylation-induced structural changes in smooth muscle myosin. Proc. Nat. Acad. Sci. USA, 2010, 107, 8207-8212.

Espinoza-Fonseca LM*, Wong C, Trujillo-Ferrara JG. Tyr74 is essential for the formation, stability and function of Plasmodium falciparum triosephosphate isomerase dimer. Arch. Biochem. Biophys. 2010, 494, 46-57.

Espinoza-Fonseca LM*. Thermodynamic aspects of coupled binding and folding of an intrinsically disordered protein: a computational alanine scanning study. Biochemistry. 2009, 48, 11332-11334.

Espinoza-Fonseca LM*. Reconciling binding mechanisms of intrinsically disordered proteins. Biochem. Biophys. Res. Commun. 2009, 382, 479-482.

Simon Gruber

Graduate Student (BMBB)

Office:	1-152 Nils Hasselmo Hall
Phone:	(612) 626-0113
Fax:	(612) 624-0632
Mail:	Dept. of Biochemistry (BMBB) University of Minnesota 6-155 Jackson Hall 321 Church St. Minneapolis, MN 55455
Email:	sjg@ddt.biochem.umn.edu

CURRICULUM VITAE

Research Interests

Ca2+ Transport and Muscle Contraction Background:

Muscle contraction is a complex process that initiates when a neuronal signal arrives at a muscle cell. Ca2+ is released from its intracellular storehouse (the sarcoplasmic reticulum, or SR), and the high [Ca2+] causes sarcomeric proteins to initiate contraction. For the muscle to relax, Ca2+ must be put back into the SR, and this is done by the sarco/endoplasmic reticulum calcium ATPase (SERCA). SERCA actively transports Ca2+ into the SR against its concentration gradient and is inhibited by phospholamban (PLB) at submicromolar [Ca2+]. This inhibition can be relieved by adrenergic stimulation, which phosphorylates PLB and alters its structural dynamics making the cytoplasmic domain's movements more rapid. A major goal of the Thomas lab is to understand how exactly PLB regulates SERCA, and how this inhibition is relieved. Our current understanding is that PLB does not dissociate from SERCA even when SERCA is fully active; instead inhibition is relieved because phosphorylated PLB (pPLB) interacts with SERCA differently than unphosphorylated PLB. Recently our lab, in collaboration with the Veglia lab at U of MN, has used spectroscopy to study the effects of PLB phosphorylation as well as various point mutations on PLB's structure and dynamics. We hypothesize that we can make mutants with dynamics similar to pPLB that will also mimic pPLB functionally.

Spectroscopic Design of Phospholamban Mutants to Treat Heart Failure:

One of the most common symptoms of heart failure (HF) is impaired calcium handling, frequently resulting from decreased SERCA activity. The recent collaborations between the Thomas and Veglia labs that have shown that PLB dynamics, measured by EPR and NMR, can be correlated with its inhibitory function, are the impetus for this project. My current goal is to co-express WT and mutant PLBs with human SERCA2a in HEK cells and measure SERCA activity, with the goal of identifying LOF-PLB mutants (PLBM) that can compete with WT-PLB and thus relieve SERCA inhibition (Fig. 1). Because displacement of WT PLB is a critical feature of PLBM, I will also try to take advantage of another HEK system being developed by Suzanne Haydon, where she is co-expressing fluorescently labeled SERCA and PLB. I plan to measure FRET between the labeled proteins in the presence and absence of PLBM, effectively measuring competition. Based on my results, rAAV will be used to test PLBM in rodent and porcine models of HF for efficacy in vivo and ability to respond to adrenergic stimulation. Data from initial trials in rodents will be used to choose the most successful mutant and expand on it, creating double or triple mutants with predictable inhibitory potency. These animal studies are critical because of the need for the mutants to compete with WT, so it is important that we demonstrate the ability to correlate our FRET measurements with in vivo competition.

Cysteine Mutagenesis to Specifically Label SERCA:

Another project I intend to begin soon is to express a “cys-lite” or cys-null SERCA in HEK cells. Cysteines are the best naturally occurring residues to label with EPR active or fluorescent probes, and SERCA has many. Currently there is only one of SERCA’s 26 cysteines that can be specifically labeled with a fluorescent probe (Cys674). The data that can be obtained from labeling of this site alone is limited, so more labeling sites are desired. This project would require mutating a minimal number of cysteines (starting with the most reactive) and introducing new ones at desired labeling sites. While no one has attempted to specifically label SERCA anywhere other than Cys674, there have been many studies on the reactivity of SERCA’s many cysteines and which mutations inhibit expression. I will use this data to remove the most reactive cysteines while not greatly reducing expression. I may also attempt to introduce tetra-cysteine labeling sites that are highly specific, and this should work without removing any cysteines. One potential problem is reduced expression after mutation of a large number of cysteines, but this will likely be overcome by the variety of approaches we are taking to the problem. Finally, I also intend to genetically tag these SERCA mutants so they may be affinity-purified and labeled.

Zach James

Graduate Student (BMBB)

Office:	5-290 Nils Hasselmo Hall
Phone:	(612) 626-3322
Fax:	(612) 624-0632
Mail:	Dept. of Biochemistry (BMBB) University of Minnesota 6-155 Jackson Hall 321 Church St. Minneapolis, MN 55455
Email:	zj@ddt.biochem.umn.edu

Research Interests:

My work focuses on phospholamban (PLB), a 52-residue membrane protein that helps govern cardiac muscle contractility by regulating the sarcoplasmic reticulum (SR) Ca-ATPase (SERCA).� Both proteins are embedded in the SR membrane of cardiomyocytes, where SERCA facilitates muscle relaxation by pumping Ca from the myoplasm into the SR lumen.� PLB binds and inhibits SERCA, reducing its apparent Ca affinity, while PLB phosphorylation at Ser16 restores Ca sensitivity without dissociating the two proteins.� The structural basis for SERCA regulation by PLB remains obscure despite several decades of research, and my goal is to elucidate this mechanism using site-directed spin labeling and electron paramagnetic resonance (EPR).

I use a variety of EPR techniques to study SERCA-PLB structural dynamics. Currently I am incorporating Cys residues into the transmembrane (TM) and cytosolic helices of PLB that serve as labeling sites for thiol-reactive spin probes. Accessibility measurements with saturation rollover EPR allow me to position the TM helix of PLB relative to the bilayer and detect any topological changes that occur upon SERCA binding and Ser16 phosphorylation. Double electron-electron resonance (DEER) experiments allow me to measure inter-spin distances within doubly-labeled PLB or between singly-labeled PLB and SERCA to detect domain motions following phosphorylation. When precise dynamics and distance measurements are needed, I use PLB constructs synthesized by Dr. Christine Karim that contain the amino acid TOAC. This spin label rigidly couples the nitroxide group to the peptide backbone and permits EPR measurements that directly reflect the structure and rotational motion of PLB.

Publications

James, ZM, JE McCaffrey, KD Torgersen, CB Karim, and DD Thomas. 2012. Oligomeric interactions in calcium transport regulation probed by electron paramagnetic resonance. Biophys J accepted August 9th 2012.

Li, J, ZM James, X Dong, CB Karim, and DD Thomas. 2012. Structural and functional dynamics of an integral membrane protein complex modulated by lipid headgroup charge. J Mol Biol 418:379-389.

Lin, AY, E Prochniewicz, ZM James, B Svensson, and DD Thomas. 2011. Large-scale opening of utrophin�s tandem CH domains upon actin binding by an induced-fit mechanism. Proc Nat Acad Sci USA 108:12729�12733.

Jillian Johnson

Undergraduate Research Assistant

Office:	1-152 Nils Hasselmo Hall
Phone:	(920) 621-6446
Fax:	(612) 624-0632
Mail:	Dept. of Biochemistry (BMBB) University of Minnesota 6-155 Jackson Hall 321 Church St. Minneapolis, MN 55455
Email:	jj@ddt.biochem.umn.edu

Research Interests:

My research focuses on the structural role of the regulatory muscle protein Myosin Binding Protein-C (MyBP-C) and its involvement in cardiac disease. I emphasize molecular biology and biochemistry methods in my project and work with techniques such as site-directed mutagenesis, expression, and purification of recombinant proteins for spectroscopic studies. I work in collaboration with postdoctoral fellow Brett Colson.

Personal:

I am originally from Green Bay, WI and am majoring in Microbiology with a Pre-Med intention. After graduation I hope to attend medical school, with the goal of eventually pursuing a career in orthopedic surgery and sports medicine. The majority of my other extracurricular involvement at the U is as a student-athlete; I compete on the women�s Track & Field and Cross Country teams as a distance runner.

Holly Langer

Junior Scientist

Office:	5-290 Nils Hasselmo Hall
Phone:	(612) 626-3322
Fax:	(612) 624-0632
Mail:	Dept. of Biochemistry (BMBB) University of Minnesota 6-155 Jackson Hall 321 Church St. Minneapolis, MN 55455
Email:	hrl@ddt.biochem.umn.edu

Ji Li

Postdoctoral Associate

Office:	5-290 Nils Hasselmo Hall
Phone:	(612) 626-3322
Fax:	(612) 624-0632
Mail:	Dept. of Biochemistry (BMBB) University of Minnesota 6-155 Jackson Hall 321 Church St. Minneapolis, MN 55455
Email:	jili@ddt.biochem.umn.edu

Research Interests

My research focus is on developing and applying biophysical spectroscopy to study membrane protein regulations, with an emphasis on interpreting biochemical regulations (function) in terms of biophysical interactions (structural dynamics). Specifically, I am using time-resolved fluorescence resonance energy transfer (TRFRET), time-resolved phosphorescence anisotropy (TPA) and fluorescence microscopy to study SERCA and its regulator PLB, two transmembrane proteins that are responsible for 70% of the Ca uptake during cardiac muscle relaxation in the human heart.

Time-resolved FRET to resolve multiple structural states:

Our lab has shown that Ca and phosphorylation relieves SERCA inhibition without dissociating PLB, but rather inducing changes of structural dynamics within SERCA or PLB. In order to further elucidate the functional regulations in terms of physical interactions, I controlled the physical interaction between SERCA and PLB using charged lipids. Then used time-resolved FRET to resolve the structural states of the SERCA-PLB complex. PLB binds to SERCA at two structural states, corresponding to two dynamic states measured using EPR. The ordered membrane associated state is more inhibitory, whereas the disordered membrane disassociated state is less inhibitory.

TPA to resolve protein aggregation:

Previous TPA work from our lab also show that PLB induces SERCA aggregation and thus reduces its ATPase activity. However, previous work were based on ErITC, a phosphorescence derivative that binds to SERCA in the ATP binding pocket, thus significantly interferes with SERCA activity. I am labeling SERCA using ErIA in the native cardiac SR vesicles, and study the effects of PLB phosphorylation on SERCA aggregation. ErIA labeled SERCA is fully functional. TPA results show that phosphorylation of PLB increases rotational dynamics of SERCA, suggesting that PLB regulates SERCA aggregation in cardiac muscles. Unphosphorylated PLB induces SERCA aggregation and thus reduced its ATPase activity. Phosphorylation of PLB decreases SERCA aggregation and thus relieves the SERCA inhibition.

Polarized TIRF to resolve domain orientation:

I developed a polarized total internal reflection fluorescence microscopy to study the orientation of PLB labeled with bi-functional fluorescence label in supported lipid bilayer. SERCA binding lifts PLB cytoplasmic domain away from the membrane surface.

I am also managing the Biophysical Spectroscopy Facility (BSF). Please go to the BSF website and check our state of the art spectroscopic instruments. We are looking forward to sharing with you our knowledge in spectroscopy, and helping you to explore the fascinating protein world.

Gage Matthews

Assistant Scientist

Office:	1-152 Nils Hasselmo Hall
Phone:	(612) 626-0113
Fax:	(612) 624-0632
Mail:	Dept. of Biochemistry (BMBB) University of Minnesota 6-155 Jackson Hall 321 Church St. Minneapolis, MN 55455
Email:	gfm@ddt.biochem.umn.edu

Jesse McCaffrey

Graduate Student (Physics)

Office:	5-290 Nils Hasselmo Hall
Phone:	(612) 626-3322
Fax:	(612) 624-0632
Mail:	Dept. of Biochemistry (BMBB) University of Minnesota 6-155 Jackson Hall 321 Church St. Minneapolis, MN 55455
Email:	jem@ddt.biochem.umn.edu

Research Interests

My primary research project is studying the dynamics of the SERCA-PLB complex using conventional and saturation transfer electron paramagnetic resonance (EPR). SERCA is an integral membrane protein located in the cellular sarcoplasmic reticulum (SR) that hydrolyzes ATP to pump calcium out of the SR, causing muscle relaxation. PLB is also an integral membrane protein, and inhibits SERCA activity upon binding. This inhibition can be reversed by phosphorylation of PLB, but the structural mechanism behind this relief of inhibition is not well understood. Elucidation of this interaction could lead to treatments for muscular defects, such as heart failure and muscular dystrophy. For example, overexpression of SERCA via AAV vectors has been shown to rescue heart failure in human clinical trials (Lipskaia et al 2010), as well as significantly increase muscle capacity in mice with muscular dystrophy (Goonasekera et al 2011).

I study the SERCA-PLB interaction using electron paramagnetic resonance. We attach spin labels to specific locations on the protein with site-directed spin labeling (SDSL) and solid phase peptide synthesis (SPPS). The EPR experiments reveal static and dynamic properties of the spin label. Using a rigid spin label such as TOAC, we can extend these spin label properties to the protein itself. To study the SERCA-PLB complex, we co-reconstitute varying amounts of SERCA and TOAC-PLB (unphosphorylated and phosphorylated) into lipid vesicles, then acquire EPR spectra to determine the rotational mobility of PLB. These deviations in rotational mobility indicate important structural changes, including conformation and protein binding. Complimentary EPR experiments, such as dipolar broadening and dipolar electron-electron resonance (DEER), produce distance measurements between adjacent spin labels, so these techniques are also implemented to provide supporting information.

My novel contributions to this project include optimization of the conditions for PLB/SERCA co-reconstitution, calibration of ST EPR as a quantitative measure, and sequential variation of protein content to demonstrate incremental effects. For simplicity, these experiments currently use a synthesized monomeric form of PLB, however we intend to probe wild type PLB, which has been shown to exist in a dynamic equilibrium between monomeric and pentameric forms. Additionally, we will explore oligomeric interactions among SERCA molecules, because this is believed to change the binding dynamics with PLB. Since electrostatic interactions are critical in the SERCA-PLB interaction (MacLennan et al 1993), we will introduce charged lipids in the co-reconstitution vesicles to study the effect on binding affinity.

Rebecca Moen

Graduate Student (BMBB)

Office:	5-290 Nils Hasselmo Hall
Phone:	(612) 626-3322
Fax:	(612) 624-0632
Mail:	Dept. of Biochemistry (BMBB) University of Minnesota 6-155 Jackson Hall 321 Church St. Minneapolis, MN 55455
Email:	rjm@ddt.biochem.umn.edu

Research Interests

My research focuses on the functional and structural perturbations of oxidative modification in muscle proteins. The production of reactive oxygen species (ROS) results from normal cellular metabolism, yet changes in protein structure and function result from excessive ROS production, with important consequences for cellular responses to environmental stressors. The accumulation of oxidatively modified proteins has been implicated in the progression of biological aging, muscle degeneration and disease. The sulfur-containing amino acids, cysteine (Cys) and methionine (Met), are the prime cellular targets of ROS. Met oxidation by ROS, and reduction by Met sulfoxide reductase, has emerged as a critical cellular process with far-reaching implications in health and disease.

My work focuses on two muscle proteins, calmodulin (CaM) and myosin (Fig. 1). CaM possesses an unusually high methionine content making it highly susceptible to oxidation. CaM Met residues are converted to methionine sulfoxide upon oxidation, impairing CaM's ability to regulate the SR Ca2+ release channel/ryanodine receptor complex (RyR). The oxidation-induced decline in CaM regulatory efficacy and the central role of CaM in RyR regulation suggests that CaM oxidation alters the structure of the channel complex, SR Ca2+ release, and ultimately muscle contraction. While considerable attention has focused on oxidative modifications of membrane proteins in muscle aging and disease, nothing is more fundamental to muscle function than force-generation itself, which is produced by myosin interacting with actin and driven by ATP hydrolysis. Oxidative stress and aging have focused on sarcomeric proteins, including myosin, and evidence is growing that myosin Met oxidation is an effective cause of functional loss due to oxidative stress in muscle. The goal is to identify and characterize Mets functionally and structurally sensitive to the redox environment. This work is crucial for gaining insight into the loss of contractile function in both skeletal and cardiac muscle. The current understanding of the underlying molecular mechanisms of Met oxidation is limited, and a much clearer picture linking specific oxidative modifications to discrete functional impacts and structural perturbations is needed.

Joe Muretta

Research Associate

Office:	5-290 Nils Hasselmo Hall
Phone:	(612) 626-3322
Fax:	(612) 624-0632
Mail:	Dept. of Biochemistry (BMBB) University of Minnesota 6-155 Jackson Hall 321 Church St. Minneapolis, MN 55455
Email:	jmm@ddt.biochem.umn.edu

Research Interests

Conformational dynamics of macromolecular assemblies in muscle: Time resolved fluorescence and EPR spectroscopy of regulated proteins.

Protein function is driven by conformational dynamics:

Nowhere is this more clear than in muscle where myosin converts the chemical energy of ATP in to mechanical work. Ligand binding and dissociation by myosin, drives structural transitions in the myosin catalytic and light-chain binding domains, which propagate nanometers from the ligand binding sites. These changes sustain pN of force, thus converting the chemical potential of ligand binding and dissociation into mechanical work. In the classic view of protein structure/function, discrete biochemical states (Ligand binding) correspond to unique structural states. This paradigm can be traced back to Perutz and the first models of hemoglobin and myoglobin. And though elegantly predictive, we know these are just snap shot images of the biomolecule. It takes dynamics to bring these images to life.

Statistical mechanics tell a more complicated story:

The conformation of protein and nucleic acids in crystals are ground state conformations, stabilized by crystallization conditions and in some cases by lattice stacking. They represent allowable conformations in the structural landscape. They do not however, reflect the state of a protein or nucleic acid in solution, much less in cells. Outside the crystal lattice, proteins are dynamic, their structures ever changing as they diffuse over the energy landscape. We need to understand the molecules entire conformational landscape if we want to understand how proteins function. Spectroscopy and molecular dynamics simulations allow us to map structural changes in proteins from the femptosecond to millisecond time scales. When combined with high resolution x-ray crystallography, these approaches allow us to see life in action.�

Myosin converts the chemical energy of ATP into mechanical work:

We use optical and magnetic spectroscopy and molecular dynamics simulations to map the conformational transitions in myosin and actin during active ATP turnover. We perform these studies in silico, in vitro, in isolated muscle fibers, and in live cells. Results from these experiments allow us to correlate structural changes in myosin and actin, with the biochemical kinetics of the actomyosin ATPase cycle. This is the key to understanding how the chemical energy of ATP is coupled to protein mechanics.

Time-resolved fluorescence:

Researchers in the Thomas lab have developed a novel high-throughput time-resolved spectrophotometer. This instrument measures a full time-resolved fluorescence decay with signal/noise of 100 or greater, in 0.1 ms. This technology opens the door to an entirely new class of experiment where time-resolved fluorescence is used to map structural changes in proteins during a biochemical transient. We call this technology "Transient time-resolved fluorescence." The time-resolved decay of a fluorescent probe is sensitive to both local and global conformational changes. Changes in the fluorescence decay, can be used to track structural transitions in proteins. We are using this approach to measure structural transitions detected by time-resolved FRET and time-resolved fluorescence anisotropy during the pre-steady state biochemical conditions of ligand binding and dissociation. Examples that I am currently working on include detecting the structural isomerization in the myosin active site associated with ATP and ADP binding. These experiments are performed with fluorescent nucleotides such as Mant-ATP and Mant-ADP and detected by transient time-resolved fluorescence anisotropy. I am also studying the structural isomerization in myosin's relay helix, converter domain, and actin binding cleft, to determine how critical biochemical events like actin binding, nucleotide binding, and phosphate release, are coordinated with structural transitions in the myosin catalytic domain.

Collaborators:

I am collaborating with members of the Thomas lab, including Christine Karim, Piyali Guhathakurta, Ewa Prochniewicz, Razvan Cornea, and Dave Thomas. I also collaborate with researchers in the Courtney Aldrich, Mark Distefano, Joe Metzger, Margaret Titus, and James Ervasti labs at the University of Minnesota as well as with researchers outside the University of Minnesota in projects using transient time-resolved fluorescence to investigate the structural dynamics of non-muscle molecular motors, and enzymes.

Publications

1. Muretta JM, Kyrychenko A, Ladokhin AS, Kast DJ, Gillispie GD, Thomas DD. High-performance time-resolved fluorescence by direct waveform recording. Rev Sci Instrum 81: 103101

2. Muretta JM, Mastick CC. 2009. How insulin regulates glucose transport in adipocytes. Vitam Horm 80: 245-86

3. Muretta JM, Romenskaia I, Cassiday PA, Mastick CC. 2007. Expression of a synapsin IIb site 1 phosphorylation mutant in 3T3-L1 adipocytes inhibits basal intracellular retention of Glut4. J Cell Sci 120: 1168-77

4. Muretta JM, Romenskaia I, Mastick CC. 2008. Insulin releases Glut4 from static storage compartments into cycling endosomes and increases the rate constant for Glut4 exocytosis. J Biol Chem 283: 311-23

5. Thomas DD, Muretta JM, Colson BC, Kast DJ. 2011. Spectroscopic probes of muscle proteins. Comprehensive Biophysics Chapter 4.15 (Accepted).

6. Wilson DJ, Shi C, Duckworth BP, Muretta JM, Manjunatha U, et al. A continuous fluorescence displacement assay for BioA: an enzyme involved in biotin biosynthesis. Anal Biochem 416: 27-38

Florentin Nitu

Postdoctoral Associate

Office:	1-150 Nils Hasselmo Hall
Phone:	(612) 626-3322
Fax:	(612) 624-0632
Mail:	Dept. of Biochemistry (BMBB) University of Minnesota 6-155 Jackson Hall 321 Church St. Minneapolis, MN 55455
Email:	fn@ddt.biochem.umn.edu

Research Interests

I am focused on a class of intracellular calcium channels, termed ryanodine receptors, or RyRs, that play a central role in controlling muscle contraction and the heart beat, more specific:

• the role of channel accessory proteins, including Calmodulin and FKBP in channel activity regulation and structure-function correlations.
• the molecular mechanisms of drugs and bioactive peptides that modulate RyR channels.
• genetically encoded fluorescent tags.

Techniques: cDNA cloning, protein mutagenesis, E. coli protein expression, recombinant protein purification, HPLC, FPLC, site-directed fluorescent labeling, fluorescence spectroscopy, CD, MS, �protein analytical chemistry.

Publications

1. Ablorh NA, Miller T, Nitu F., Gruber SJ, Karim C, Thomas DD. Accurate quantitation of phospholamban phosphorylation by immunoblot. Anal Biochem., 2012, 425(1):68-75.
2. Guo T., Fruen B.R., Nitu F.R., Nguyen T.D., Yang Y., Cornea R.L., Bers D.M. FRET Detection of Calmodulin Binding to the Cardiac RyR2 Calcium Release Channel, Biophys. J., 2011, 101(9):2170-7.
3. Cornea R.L., Nitu F.R., Sams� M., Thomas D.D., Fruen B.R. Mapping the ryanodine receptor FK506-binding protein subunit using fluorescence resonance energy transfer. J. Biol. Chem., 2010, Jun 18; 285(25):19219-26.
4. Ablorh NA, Nitu F., Engebretsen K, Thomas DD, Holger JS. Insulin-dependent rescue from cardiogenic shock is not mediated by phospholamban phosphorylation. Clin. Toxicol. (Phila). 2009 Apr; 47(4):296-302.
5. Cornea R.L., Nitu F., Gruber S., Kohler K., Satzer M., Thomas D.D., Fruen B.R. FRET-based mapping of calmodulin bound to the RyR1 Ca2+ release channel. Proc. Natl. Acad. Sci. U S A, 2009, Apr 14; 106(15):6128-33.
6. Zamoon J., Nitu F., Karim C., Thomas D.D., Veglia G. Mapping the interaction surface of a membrane protein: unveiling the conformational switch of phospholamban in calcium pump regulation. Proc. Natl. Acad. Sci. U S A, 2005 Mar 29; 102(13):4747-52.
7. Fruen B.R., Balog E.M., Schafer J., Nitu F.R., Thomas D.D., Cornea R.L. Direct detection 7/8/2011of calmodulin tuning by ryanodine receptor channel targets using a Ca2+-sensitive acrylodan-labeled calmodulin. Biochemistry, 2005, Jan 11; 44(1):278-84.

Abstracts and Posters

�Florentin R. Nitu, Razvan L. Cornea, David D. Thomas, Bradley R. Fruen. University of Minnesota, Minneapolis, MN, USA. Direct detection of domain peptide binding to the cardiac ryanodine receptor (RYR2) using FRET. Biophysical Society 55th Annual Meeting, March 5-9, 2011, Baltimore, Maryland, USA.

� Bradley R. Fruen, Mallory Turner, Florentin R. Nitu, David D. Thomas, Razvan L. Cornea. University of Minnesota, Minneapolis, MN, USA. FRET detection of Calmodulin binding and structural rearrangements within the cardiac RyR2 calcium release channel (1567-Pos). Biophysical Society 54th Annual Meeting, February 20 � 24, 2010, San Francisco, California, USA.

� Razvan L. Cornea, Florentin R. Nitu, David D. Thomas, Bradley R. Fruen. University of Minnesota, Minneapolis, MN, USA. Time-Resolved FRET detection of structural distributions involving FKBP12.6 and Calmodulin bound within macromolecular RyR channels (1558-Pos). Biophysical Society 54th Annual Meeting, February 20 � 24, 2010, San Francisco, California, USA.

� Razvan L. Cornea, Florentin Nitu, Katherine Kohler, David D. Thomas, Bradley R. Fruen. Structural Characterization of FKBP Interactions with RyR Channels Using Site-Directed Fluorescent Labeling and FRET. Biophysical Society 53th Annual Meeting, February 28 � March 4, 2009, Boston, Massachusetts, USA.

� Bradley Fruen, Florentin Nitu, David Th omas, Razvan Cornea. FRET-based mapping of calmodulin bound to ryanodine receptor channels. Calmodulin Modulation of Ion Channels Asilomar Conference Center, October 30 - November 2, 2008, Asilomar, California, USA.

� Engebretsen KM, Holger JS, Harris CR, Ablorh N, Nitu F, Thomas D. Therapeutic Misadventure of High Dose Insulin without Adverse Effects. Clinical Toxicology 2008; 46(7):604. Presented at North American Congress of Clinical Toxicology, Toronto, Ontario, CA, September 2008. 

� Razvan L. Cornea, Florentin R. Nitu, Michael B. Satzer, David D. Thomas, Bradley R. Fruen. University of Minnesota, Minneapolis, MN, USA. Fluorescent probe of FKBP interactions with ryanodine receptor channels, Biophysical Society 51th Annual Meeting, March 3-7, 2007, Baltimore, Maryland, USA.

� Bradley R. Fruen1, E.M. Balog1, Ruiwu Wang2, S.R.W. Chen2, F. Nitu1, R.L. Cornea1. 1University of Minnesota, Minneapolis, MN, USA, 2University of Calgary, Calgary, AB, Canada.� Calmodulin regulation of the type 3 ryanodine receptor , Biophysical Society 49th Annual Meeting, Presentation Session: (421-Pos/B260)� , February 12-16, 2005, Long Beach, California, USA.

Kris O'Meara

Undergraduate Office Assistant

Phone:	(507) 319-3745
Email:	omear048@umn.edu

Lab Duties

Dishwashing, collecting and delivering departments mail, managing departments calendar, solution and media preparation, general lab maintenance.

Karl Petersen

Graduate Student (BMBB)

Research Interests

My research is focused on skeletal muscle myosin and its interaction with actin filaments. As an undergraduate, I do most work in collaboration with Dr. Joseph M. Muretta. We use an in vitro model of muscle with myosin harvested from the amoeba Dictyostelium discoideum and actin harvested from rabbit skeletal muscle. To investigate structural changes in myosin, we exploit structural models to design labeling sites which are then created by site-directed mutagenesis of the myosin gene. Once purified, this myosin is labeled with various fluorescent probes. We then use FRET to measure distances within myosin during its enzymatic cycle. Current work uses a pair of probes on a structure called the relay helix. This helix bends during the myosin powerstroke, causing the distance between probes to decrease.

We combine this structural information with kinetics to generate kinetic models describing muscle contraction at the molecular level. To further develop the structural kinetics of myosin, we use phosphate-binding protein (PBP), a protein secreted by E. coli. When labeled with a fluorescent probe, PBP becomes an effective sensor for phosphate release at rates seen during the myosin enzymatic cycle (~20/s). We are also able to measure the rate of myosin binding to actin by labeling actin with pyrene. In this way we measure three events during the myosin powerstroke: relay helix bending, phosphate release and relay helix bending.

In support of this work on myosin, we also develop technology to acquire and process fluorescence data from the "Fargoland" transient time-resolved spectrometer. This (TR)2 instrument uses a pulsed laser to measure fluorescence decays on the ns time scale. By integrating these decays, a ms scale measurement is obtained allowing us to compare our novel (TR)2 instrument with conventional instruments. In addition, we are developing methods for use with the ns scale microplate-based spectrometer "NovaFluor" (Fluorescence Innovations, Inc.). The microplate format is exciting because it will permit us to map any experiment onto a 2-dimensional concentration gradient, potentially accelerating discovery.

Allison Raney

Undergraduate Research Assistant

Lab Duties

Dishwashing, solution and media preparation, general lab maintenance.

Mike Schaid

Undergraduate Volunteer

Research Interests

My research is focused on the muscle proteins sarcolipin and phospholamban. I am working with Mike Autry, a postdoc in the Thomas Lab. Much of my work involves investigating the potential for these proteins to self-assemble into ion channels. Currently, we are investigating channel potential using an E. coli expression system. I am also helping Dr. Autry examine sarcolipin and phospholamban interactions with SERCA. We use an insect cell-baculovirus expression system to harvest protein for further analysis with fluorescence microscopy.

Bengt Svensson

Research Associate

Office:	5-126 Nils Hasselmo Hall
Phone:	(612) 626-3225
Fax:	(612) 624-0632
Mail:	Dept. of Biochemistry (BMBB) University of Minnesota 6-155 Jackson Hall 321 Church St. Minneapolis, MN 55455
Email:	svens005@umn.edu

Research Interests

My research is part of the larger effort in the David D. Thomas lab on the study of structural dynamics associated with force-generating proteins during muscle contraction and the molecular mechanisms of catalysis and regulation of the Ca-ATPase (SERCA) that pumps calcium into the sarcoplasmic reticulum and thus relaxes the muscle. My goal is specifically, to use molecular modeling and molecular dynamics simulations to aid the interpretation of experimental results and to generate new structural and dynamical models that describe the function of muscle proteins on an atomistic level.

I work on many different projects, the main project currently is to use molecular dynamics simulations on SERCA labeled with the fluorescent probe IAEDANS with the aim to establish a more rigorous foundation for analyzing spectroscopic data. Site-specific labeling of a protein with a fluorescent probe can provide insight into local structural dynamics, based on fluorescence quenching or anisotropy measurements, or used to measure distances based on fluorescence resonance energy transfer (FRET) to another label. SERCA was labeled at position Cys674 in the P-domain (shown blue in figure) with the fluorescent probe IAEDANS. The starting point for the molecular dynamics simulations of the fluorescent probe and its protein environment was a new crystal structure of IAEDANS-labeled SERCA that was determined in collaboration with Howard Young, Univ. of Alberta, Edmonton. The structure was determined to 3.4� resolution, which was sufficient to show the IAEDANS label in close proximity to residues Arg615 and Arg620. To be able to perform these simulations, we developed CHARMM force-field parameters for the fluorescence probe IAEDANS. Quantum chemistry calculations have also been performed on the ground state and excited states of IAEDANS, to determine the orientation of the transition dipole moment. The transition dipole autocorrelation functions and reorientation times were calculated from the simulated trajectories and compared with experimental measurements by fluorescence anisotropy. FRET parameters were also determined from the simulations and compared with results from fluorescence experiments using IAEDANS as the donor and TNP-ADP bound in the nucleotide pocket as the acceptor. The results show that we have established a reliable framework for both fluorescence experiments and MD simulations in this system.

Another project is the simulation of XFP-SERCA fusion proteins to compare with experimental data from FRET. This project is a collaboration with Seth Robia, Loyolla University, Chicago. An ensemble of conformations of the XFP fused to SERCA is generated using a Monte-Carlo based method. From that ensemble of structures we can calculate FRET parameters using both distance distributions and orientation information and compare that to the experimental data. The aim is to understand the dynamics i.e. conformational changes in the cytoplasmic domains in SERCA.

Support:

My current work has been supported by NIH (GM27906, AR007612) and the Minnesota Supercomputing Institute.

Research Experience:

I have mostly been doing theoretical research on protein structure and function, but have significant laboratory experience as well in the field of biophysics/spectroscopy and biochemistry. Before the current work on muscle proteins I have worked on the opioid receptors and GPCR's, and on the photosynthetic proteins involved in oxygen evolution i.e. photosystem II.

The theoretical techniques I have experience with include molecular modeling of proteins, molecular dynamics simulations, quantum chemistry calculations of protein ligands and spectroscopic probes. I have also used bioinformatics tools to identify important residues and regions of protein sequences based on multiple sequence alignments.

The experimental measurement techniques I have used include EPR, FTIR, CD, fluorescence, UV/Vis, thermoluminescence, SAXS, mass-spectroscopy, steady state and flash oxygen polarography etc. I was member of a team involved in the development of kinetic fluorescence and oxygen polarography instruments at UIUC. The instrument we designed were commercially available through Artisan Scientific, Champaign, IL

Software I use include, CHARMM, CHARMM-GUI, MMTSB toolset, VMD, DS Visualizer, OriginPro, ProFit, CE (Combinatorial Extension), Gaussian/Gaussview, Molden, ConQuest, Situs, FPMOD, pyMol, DOWSER, Reduce, Procheck, ClustalW, EMBOSS.

Other Skills:

System administration experience on Linux (including setting up a small HPC cluster), UNIX (SGI), and Microsoft Windows environments. Computer hardware maintenance, repairs, and upgrades. Programming in Fortran, C, and scripting languages Perl, C-shell. User interface programming in Perl/Tk.

Education:

B.S. Chemistry & Microbiology, Stockholm University, Sweden, 1990
Ph.D. Biochemistry, Stockholm University, Sweden (S. Styring) 1996

Post Doc:

Center for Biophysics and Computational Biology and Dept. of Microbiology, University of Illinois (A. Crofts) 1996-99
Dept. of Biochemistry, Molecular Biology and Biophysics, University of Minnesota (B. Barry) 1999-02
Dept. of Medicinal Chemistry, University of Minnesota (D. Ferguson) 2002-07
Dept. of Biochemistry, Molecular Biology and Biophysics, University of Minnesota (D.D. Thomas) 2007-present

Recent Publications:

19. Lin, A. Y., E. Prochniewicz, Z. M. James, B. Svensson, and D. D. Thomas (2011) "Large-scale opening of utrophin's tandem CH domains upon actin binding, by an induced-fit mechanism" Proc. Natl. Acad. Sci. U.S.A., 108: 12729�12733.
18. Rose, S., Minagawa, J., Seufferheld, M., Padden, S., Svensson, B., Kolling, D.R.J., Crofts, A.R. and Govindjee (2008) "D1-arginine257 mutants (R257E, K, and Q) of Chlamydomonas reinhardtii have a lowered QB redox potential: analysis of thermoluminescence and fluorescence measurements", Photosynth. Res. 98, 449�468.
17. Klein, J.C., Burr, A.R., Svensson, B., Kennedy, D.J., Allingham, J., Titus, M.A., Rayment, I. and Thomas, D.D. (2008) "Actin-binding cleft closure in myosin II probed by site-directed spin labeling and pulsed EPR", Proc. Natl. Acad. Sci. U.S.A. 105, 12867-12872.
16. Winters, D.L., Autry, J.M., Svensson, B., and Thomas, D.D. (2008) "Interdomain fluorescence resonance energy transfer in SERCA probed by cyan-fluorescent protein fused to the actuator domain", Biochemistry 47, 4246-4256.
15. Goodell, J.R., Svensson, B., and Ferguson, D.M. (2006) "Spectrophotometric Determination and Computational Evaluation of the Rates of hydrolysis of 9-Amino-Substituted Acridines", J. Chem. Inf. Model. 46, 876-883.

Corrin Laposki

Undergraduate Ressearch Assistant

Office:	1-152 Nils Hasselmo Hall
Phone:	(612) 626-0113
Fax:	(612) 624-0632
Mail:	Dept. of Biochemistry (BMBB) University of Minnesota 6-155 Jackson Hall 321 Church St. Minneapolis, MN 55455
Email:	ckl@ddt.biochem.umn.edu

Matt Mauseth

Graduate Student

Office:	1-152 Nils Hasselmo Hall
Phone:	(612) 626-0113
Fax:	(612) 624-0632
Mail:	Dept. of Biochemistry (BMBB) University of Minnesota 6-155 Jackson Hall 321 Church St. Minneapolis, MN 55455
Email:	mm@ddt.biochem.umn.edu

Tory Schaaf

Graduate Student

Office:	1-152 Nils Hasselmo Hall
Phone:	(612) 626-0113
Fax:	(612) 624-0632
Mail:	Dept. of Biochemistry (BMBB) University of Minnesota 6-155 Jackson Hall 321 Church St. Minneapolis, MN 55455
Email:	tms@ddt.biochem.umn.edu