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Department of Chemistry and Chemical Biology at Rensselaer Chemistry and Chemical Biology
Richard Gross
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Richard Gross

Constellation Professor of Biocatalysis and Metabolic Engineering
Professor, Department of Chemistry and Chemical Biology
Professor, Department of Biomedical Engineering

Education:

UMASS Amherst (Prof. Robert Lenz)
BioMacromolecules
1986-1988

Polytechnic University (Prof. Mark Green)
Chemistry
Ph.D. 1986

SUNY Albany
Chemistry
B.S. 1979

Apppintments:

7/1/2013-to present
RPI: Center for Biotechnology and Interdisciplinary Studies, Constellation Chair; Professor, Dept. of Chemistry and Biology; Professor Department of Biomedical Engineering.  

1998-6/30/13
Herman F. Mark Chair, NYU-POLY

2008-present
Chief Technical Officer, SyntheZyme

2000-present
Director - NSF Center for Biocatalysis and Bioprocessing of Macromol.

2001-2007
Director - NYU-POLY Biomedical Engineering Program

2003-2006
Director: Polymer Research Institute (PRI)

2002-2005
Engineering Conferences Foundation, Board of Directors

1999
President- Biodegradable Polymer Society

1993-1998
Co-Director: NSF Center for Biodegradable Polymer Research

1988-1998
University of Massachusetts Lowell (Assistant Professor, 88-91; Associate Professor, 92-95; Full Professor, 96-98)

Honors

NSF: Presidential Young Investigator Award (1990-1995)

NSF-EPA: Presidential Award in Green Chemistry (2003)

Turner Alfrey Visiting Professorship (2010)

Johnson and Johnson: Focused Giving Award (2000-2002)

Editorial Board Member: J. Bioactive and Biocompatible Polymers (2001-present); Biomacromolecules (2000-present); Industrial Biotechnology (2005 to present); Journal of Molecular Catalysis B: Enzymatic (2005-present); Enzyme and Microbial Technology (EMT) (2005-present)

Research Areas:

Overview:

Biocatalysis encompasses a wide platform of chemistries that afford opportunities for innovative new products and processes. Hallmarks of biocatalysts are their ability to operate under mild conditions, with impressive selectivity, on a diverse range of natural and non-natural substrates. Rapid advances in biotechnology continues to decrease the time and resources required to engineer organisms to produce desired products in high titers as well as to engineer enzymes with increased thermal stability, efficiency and specificity.  Our group works with collaborators to apply existing, or develop new, biocatalytic methods that solve short or long term chemistry challenges.

With sufficient focus on structure-property relationships of natural and biobased materials, they can become next-generation replacements for a wide array of currently used non-sustainable petroleum-derived materials.  While most attention is focused on biomass for transportation fuels, less is paid to biomass as a source for organic chemicals including polymeric materials.  This is true even though biomass-derived carbon can realistically replace most or possibly all nonfuel chemical uses, which comprise ~13% of the crude oil consumed by the U.S. today. Our program explores a wide range of opportunities for use in biobased replacements such as biobased and degradable polymers, biosurfactants, biofibers for commposite reinforcement and advanced materials opportunities such as in photovoltaic materials for energy harvesting; composite materials used, for example, in wind-blade turbines for wind energy harvesting, epoxies for circuit boards and much more.  

Summary of Current Research Interests

Applying skills in biocatalysis and organic/polymer chemistry to the synthesis of biobased polymers, surfactants and peptides; reactive processing; applications in material science, antimicrobials, colloid systems, self-assembly and photovoltaics; bioresorbable functional polymeric materials for regenerative medicine; new therapeutics from glycolipids; biocatalytic reactions using whole cell and cell-free systems; enzyme structure-activity studies; biofibers; nanocomposites; designing sustainable processes; fermentation engineering. 

Developing new monomers from yeast

In preparation for weaning ourselves from a petrochemical society in which the infrastructure for extraction of energy from fossil fuels effectively subsidizes the production of plastics, it is important to develop alternatives that can be synthesized from renewable materials without compromising performance. Our laboratory is actively exploring routes to biobased plastics by combining tools of biotechnology and green chemical methods.  Examples of two monomers that are first produced by yeast fermentation and subsequently used as building blocks for polymer synthesis are described below. 

Cutinases

Increasingly, scientists are being challenged to solve environmental problems due to chemical processes that generate toxic by-products, require high energy consumption, and use petroleum based feedstocks. There is an urgent need to develop efficient biocatalytic processes that provide solutions to these problems. In this program we address these concerns by focusing on a promising family of enzymes known as cutinases that, thus far, have received disproportionally little attention relative to other ester hydrolase enzyme families. This program is organized to gain fundamental information on how the cutinase from Aspergillus oryzae (AoC) can be rationally designed and functionally characterized to improve its thermal and pH stability as well as its activity for important biocatalytic transformations such as: i) surface modification of industrially important materials to tailor their physico-chemical properties by mild-processes ii) improving the efficiency at which cutins in plant biomass are converted to chemical feedstocks and iii) developing enzymatic routes to recycle polyesters such as PET that were not designed to be degraded by enzymes. 

The natural function of cutinases is to degrade cutin, an insoluble, crosslinked, lipid-polyester matrix comprised of C16 and C18 w-hydroxy and epoxy fatty acids. Consequently, cutinases are secreted by fungal phytopathogens to modify cutin at plant surfaces to allow pathogens entrance to plants. The 0.1-to-10 mm outer membrane of higher plants is a layered composite of lipids (a waxy waterproofing) and the primarily aliphatic insoluble polyester cutin (the structural support). Remarkably, the plant cuticular barrier outperforms many engineered polymer films of comparable thickness as well as resisting the penetration of pesticides used in crop protection and foreign molecules from the atmosphere. Cutin composition differs depending upon the plant species.  Large variations in monomer composition have been reported that are specific to each cutin. 

In this program we are determining the critical structural variables of AoC and other related cutinases that will enhance its effectiveness in surface modification or hydrolysis of polyesters. Indeed, enzymes have been successfully used for many years for textile cleaning. Most prominent examples are cellulases used to reduce pilling of cotton fabrics and lipases in detergent formulations for removal of oil or grease residues. An inherent advantage of enzymes in surface modification is that their action is restricted to polymer surfaces. This is due to the large size of proteins that are generally immiscible with synthetic polymer materials. Mild conditions of enzyme-catalysis translate to milder treatment options that cause less damage to the bulk of materials and reduce energy consumption. 

w-Hydroxyfatty acids

Poly(ω-hydroxyfatty acids) have the potential to perform as functionally equivalent to versatile plastics like polyethylene while providing other attributes. The monomers required to make these polymers are also valuable in chemical products such as lubricants, adhesives, cosmetic ingredients, and anticancer therapeutics. Most work on bioplastics from renewable sources has focused on polyhydroxyalkanoates and polylactic acid. However, these materials suffer from significant performance deficits that have created difficult challenges to their adoption as general replacements for petroleum-based plastics. Poly(ω-hydroxyfatty acids) appear to overcome the functional limitations of other bioplastics, but the monomers have not been available with economics required for their conversion to commodity plastics

C. tropicalis is an attractive organism for biotransformations: it grows robustly and tolerates high concentrations of substrates which it takes up readily. In collaboration with DNA2.0, we developed a strain of C. tropicalis capable of producing high levels of ω-hydroxyfatty acids by engineering a 90% reduction in the activity of the endogenous pathways that normally metabolize these compounds to diacids. The strain engineering was achieved by identifying target sequences using PCR with degenerate primers based on known C. tropicalis and C. albicans genes, then eliminating these genes with a 2-step genome modification method that allows indefinite reuse of the same selective marker.

Future work will address opportunities to identify wild type or create engineered P450’s that allow efficient w-hydroxylation of a broad range of fatty acids that differ widely in chain length and unsaturation. Furthermore, additional strain engineering will permit a broader range of biochemical products for use as monomer building blocks, surfactants, nutrition and much more.

Protease catalyzed oligopeptides

Peptides are functionally rich, and their use in materials with properties such as pH-sensitivity, self-assembly, bioresorbability and bioactivity are areas of intense interest. Often, peptide-based materials are based on amino acid sequences found in important natural proteins such as collagen, keratin, elastins and silk.  Conventional synthesis of involves either solid phase or liquid phase peptide synthesis. These synthetic methods provide peptides in high purity with precise sequence and chain length. However, both solid phase and liquid phase peptide synthesis is costly, uses toxic reagents, requires protection-deprotection chemistry and product purification. Fermentative routes to peptides are also important but generally suffer from low product yields and are inherently difficult to adapt towards incorporation of non-peptidic end-group moieties. To increase the viability of using peptides in an ever expanding range of exciting applications, new methods for peptide synthesis are needed that are safe, scalable, and cost-effective.

When a single product with a precise amino acid sequence is not required, there is an opportunity to introduce new peptide synthetic methodologies that are simple, cost-effective and environmentally friendly. For this purpose, our laboratory is developing protease-catalyzed routes to peptides from amino acid alkyl esters.

Biobased Surfactants: The world surfactant industry depends on chemical surfactants for a wide range of applications within household, environmental, industrial and medical sectors. The world surfactants market reached US $24.33 billion in 2009, up nearly 2% from the previous year. By 2018, it is anticipated that the global surfactant market will generate revenues of more than US $41 billion, which corresponds to an average annual growth of 4.5%. Hence, the demand for surfactants continues to rise based on general growth in the chemical sector and new applications in which surfactants are key ingredients. The major industries that rely on surfactants are industrial cleaners (9%), body care and cosmetics (9.5%) and pulp and paper (11%). Other industrial products that also depend on surfactants include agrochemicals, photo chemicals, oil field chemicals, construction materials, foodstuffs, adhesives, lubricants, metal working and mining. Surfactants are used as detergents, wetting agents, emulsifiers, foaming agents, dispersants and more. Primary goals driving new innovations in surfactant technologies are to reduce surfactant toxicity, develop more efficiently ‘green’ synthetic routes for their manufacture, enhance biodegradability, increase their biobased carbon content and expand the family of surfactants that can be safely used in medical, agricultural, environmental and food applications

Biobased surfactants, produced by the chemical conjugation of biobased building blocks, constitute only 25% of total surfactant production. This is largely due to cost-performance measures of biobased surfactants that are not competitive with current petroleum based products. The goal of our research in biosurfactants is to address the opportunity to develop biobased surfactants synthesized by ‘green’ synthetic routes that, on a cost-performance basis, are competitive with their petroleum-derived counterparts. To achieve this goal we use a highly efficient fermentation process that converts sugars and plant oils to glycolipids known as sophorolipids (SLs). Modification of SLs by chemical and enzymatic methods is used to create a library of modified sophorolipids that will are being evaluated to determine their performance attributes relative to current competitive products. Strategies for SL modification focus on methods that are simple and green so that they have the potential to be used commercially. Property studies of modified sophorolipids include: i) studying their basic interfacial behavior with various oil phases, ii) determining their antimicrobial activity on organisms such a plant and human pathogens, iii) investigating their ability to self-assemble into various nano- and microstructures. 

Bioresorbable Polymers:

It is critically important that simple methods are developed to prepare functional bioresorbable polymers for a myriad of applications in polymer therapeutics. Functionality in these materials enables the introduction of bioactive moieties that can be used for targeting during drug delivery, to direct differentiation of stem cells in 3-D matrices, form complexes with DNA or SiRNA for gene therapy and to control bioresorption rates by varying the polymers affinity to water. Natural polymers such as polysaccharides and proteins are an important family of functional bioresorbable polymers that offer potential advantages such as biological recognition. However, natural polymers have drawbacks such as difficulties in purification, complex structures, immunogenicity and poor thermal stability. Generally, the latter prohibits processing of these polymeric materials by conventional melt extrusion and injection molding. In contrast, synthetically derived polymers enable the preparation of highly variable structures with a wide range of properties and functional groups. The purities of monomers and polymers are readily controlled eliminating concerns associated with natural polymer such as contamination with endotoxins. 

Our work in developing functional bioresorbable polymers combines the disciplines of biocatalysis, conventional chemistry and biobased building blocks. In on approach, a functional glycolipid known as sophorolipids (SLs) is produced by fermentation of a yeast microorganism. Lactonic SLs from this fermentation consist of the disaccharide sophorose and a hydroxyl lipid (normally, 17-hydroxyoleic acid). The fact that lactonic SLs have a double bond within their structure allows their metathesis-catalyzed ring-opening polymerization (ROP). By this approach we translate the complexity from natural SL building blocks to a polymer structure that is unique, has bonds that allow its hydrolytic degradation, and functionality that can be modified to introduce a variety of bioactive moieties. 

Poly(SL) is a solid at room temperature that undergoes a glass transition at 61 °C and melts at 123 °C. The crystal phase is associated with ordered packing of aliphatic chain segments. Semicrystalline poly(SL) also displays a long-range order (d = 2.44 nm) involving sophorose groups that is found to persist after crystal phase melting (in high-T diffractograms) with a slightly shortened distance (2.27 nm). Upon annealing at 80 °C, poly(SL) recrystallizes and, concomitantly, the disaccharide units space out again at 2.44 nm. An exothermal phenomenon that immediately follows melting and is revealed by TMDSC might be associated with the observed adjustment of sophorose units spacing in the melt.

Currently, work is in progress to create porous scaffolds by various means such as by electro-spinning (in collaboration with Professor Scandola at the University of Bologna), thermal processing with salts or other polymers that can be removed by dissolution in water, and then characterizing the polymer biocompatibility in the presence of various cell types as well as its potential bioactivity to steer cell differentiation. Also, methods for the attachment of various bioactive moieties such as peptides to polysophorolipids are under study. In addition studies of the solid state properties of poly(sophorolipids) as a function of varying the lipid and sophorose head group structure is underway.

Writing with enzymes “enzyme ink”

Our laboratory is developing methods for the selective deposition of enzymes to define high resolution features on biocompatible polymer substrates without adversely affecting the polymer substrates biocompatibility. Thus far, research aimed at ‘writing with enzymes’ has focused on highly sophisticated and expensive technologies to deliver the enzymes. For example, AFM tips modified with enzymes have been used to create surface features. Although these methods can generate high-resolution patterns, they have surface area limitations due to the capacity of AFM devices. Moreover, the depth of features is severely limited as the biocatalyst is immobilized and cannot diffuse beyond the surface of the film.

A second approach to using enzymes to lithographically define polymer films is micro-contact printing (µCP).  In this approach, enzymes are typically immobilized on the surface of polymeric stamps. These stamps, when placed in contact with the polymer surfaces, induce chemical reactions to locally degrade the polymer thereby creating patterns. Chemical immobilization of the enzyme that writes on the surface was believed to be important in order to limit the enzyme’s lateral diffusion that would lower pattern resolution.  However, immobilization of enzymes on the stamp surface imposes severe limitations on the throughput of the process. That is, the stamp must be held in contact against the surface for the time required to catalytically decompose the material to create the desired pattern. Consequently, this limits the thickness of the layer that can be defined.

Our work is exploring alternative techniques for selectively transferring an enzyme onto a polymer surface to catalyze polymer degradation thereby creating lithographically defined patterns in the polymer film.  For example, studies have been conducted to create well defined patterns within relatively thick polymer films (300-500 nm thick). Such patterned features can be used for fabrication of devices including micro-wells that can contain and release biologically active molecules in a biocompatible environment.  Current work is using poly(ε-caprolactone), PCL, films as polymer substrates and enzymes from the lipase and cutinase families. We believe that surface patterning of biomaterials is a powerful approach to control interactions with biological systems or to deliver medications that may be specifically located in patterned regions. Moreover, the use of enzymes to write on PCL via hydrolysis reactions results in water-soluble degradation products, which are safer (i.e. more biocompatible) than lithographic inks.

Lipase-catalyzed Condensation Polymerization:

Traditional chemical catalysts for polyester synthesis have enabled the generation of important commercial products. Undesirable characteristics of chemically catalyzed condensation polymerizations include the need to conduct reactions at high temperatures (150–280 °C) with metal catalysts that are toxic and lack selectivity. The latter is limiting when aspiring towards synthesis of increasingly complex and well-defined polyesters. Our laboratory has been developing methods and new functional polymeric materials that use immobilized enzyme-catalysts for condensation polyester synthesis. Unlike chemical catalysts, enzymes function under mild conditions (≤100 °C), which enables structure retention when polymerizing unstable monomers, circumvents the introduction of metals, and also provides selectivity that avoids protection–deprotection steps and presents unique options for structural control.

An important example is use of enzyme-regioselectivity to polymerized multifunctional monomers (functionality ≥3) to linear or near-linear homo- and copolymers. Our laboratory reported copolymerizations with polar multifunctional polyols that were performed under bulk reaction conditions, without activation of carboxylic acids. To avoid the use of deactivating polar-aprotic solvents, polyols were combined with monomers to form monophasic liquids at temperatures sufficiently low to maintain immobilized CALB (N435) activity (≤95 °C). For example, N435-catalyzed bulk polycondensations were implemented at 70 °C under vacuum (40–60 mmHg) using adipic acid (A), 1,8-octanediol (O) and glycerol (G) as co-monomers [monomer feed ratio (A:O:G) 1:0.8:0.2 mol/mol]. Initially, the reaction media was a two-phase liquid; however, within 60 min, the media became monophasic with suspended N435. Products at 45 min and at 2 h had little or no unreacted monomers as well as low Mn and Mw/Mn. Extension of polycondensations to 6 h and 18 h resulted in substantial increases in Mn as well as a broadening of the molecular weight distribution. Furthermore, CALB regioselectivity circumvented branching during chain formation for polymerizations up to 18 h. More recently we prepared a unique family of branched polytriglycerides by N435-condensation catalyzed polymerization of fatty acid derived diacids, glycerol and unsaturated fatty acids. 

Current challenges that we are addressing to advance the field of enzyme-catalyzed condensation are the following: i) to increase enzyme activity such that lower catalyst quantities can be used in reactions, ii) increase enzyme thermal stability to enable reactions to be conducted at higher temperatures such as above 110 oC to decrease diffusion contraints in viscous reaction media and iii) to develop immobilized enzyme systems that retain high catalytic activity but allow the re-use of enzymes over multiple reaction cycles and iv) create enzyme catalysts that provide improved activity on substrates that currently have low reactivity with available enzyme systems.

Selected References:

Ganesh, Manoj; Nachman, Jonathan; Mao, Zhantong; Lyons, Alan; Rafailovich, Miriam; Gross, Richard A. “Patterned Enzymatic Degradation of Poly(ε-​caprolactone) by High-​Affinity Microcontact Printing and Polymer Pen LithographyBiomacromolecules  14(8), 2470-2476 (2013).  

Xie, Wenchun; Teraoka, Iwao; Gross, Richard A. “Reversed phase ion-pairing chromatography of an oligolysine mixture in different mobile phases: Effort of searching critical chromatography conditions”  Journal of Chromatography A 1304, 127-132 (2013).  

Celli, Annamaria; Marchese, Paola; Sullalti, Simone; Cai, Jiali; Gross, Richard A. “Aliphatic/aromatic copolyesters containing biobased ω-hydroxyfatty acids: Synthesis and structure-property relationships” Polymer, 54(15), 3774-3783 (2013).  

Peng, Yifeng; Decatur, John; Meier, Michael A. R. and Gross, Richard A. “Ring-Opening Metathesis Polymerization of a Naturally Derived Macrocyclic Glycolipid, Macromolecules, 46(9), 3293-3300 (2013).  

 Qin, Xu; Wenchun, Xie; Tian, Sai; Yuan, Han; Yu, Zheng; Butterfoss, Glenn L.; Khuong, Anne C.; Gross, Richard A. “Enzyme-Triggered Hydrogelation via Self-Assembly of Alternating Peptides”, Chem. Commun., 49(42), 4839 – 4841 (2013).

Zhang, Yu-Rong; Spinella, Stephen; Xie, Wenchun; Cai, Jiali; Yang, Yixin; Wang, Yu-Zhong; Gross, Richard A. “Polymeric triglyceride analogs prepared by enzyme-catalyzed condensation polymerization” European Polymer Journal,  49(4), 793-803 (2013).

Zhu, Jianhui; Cai, Jiali; Xie, Wenchun; Chen, Pin-Hsuan; Gazzano, Massimo; Scandola, Mariastella; Gross, Richard A. “Poly(butylene 2,5-furan dicarboxylate), a Biobased Alternative to PBT: Synthesis, Physical Properties, and Crystal Structure” Macromolecules 46(3), 796-804 (2013).

Qin, Xu; Khuong, Anne C.; Yu, Zheng; Du, Wenzhe; Decatur, John; Gross, Richard A. “Simplifying   alternating peptide synthesis by protease-catalyzed dipeptide oligomerization” Chemical Communications  49(4), 385-387 (2013).

Bhangale, Atul S.; Beers, Kathryn L.; Gross, Richard A.; “Enzyme-catalyzed polymerizations of end-functionalized polymers in a microreactor” Macromolecules 45:7000−7008 (2012

Ganesh, Manoj; Gross, Richard A. Enzymatic biomaterial degradation: Flow conditions & relative humidity, Polymer, 53:3454-3461 (2012)

Viswanathan, Kodandaraman; Schofield, Mark H.; Teraoka, Iwao; Gross, Richard A. Surprising metal binding properties of phytochelatin-like peptides prepared by protease-catalysis, Green Chemistry14, 1020–1029 (2012)

Baker, Peter James; Poultney, Christopher; Liu, Zhiqiang; Gross, Richard; Montclare, Jin Kim Identification and comparison of cutinases for synthetic polyester degradation, Appl Microbiol Biotechnol 93:229–240 (2012)

Liu, Chen; Liu, Fei; Cai, Jiali; Xie, Wenchun; Long, Timothy E.; Turner, S. Richard; Lyons, Alan; Gross,  Richard A. Polymers from Fatty Acids: Poly(ω-hydroxyl tetradecanoic acid) Synthesis and Physico-Mechanical Studies Biomacromolecules, 12(9), 3291-3298 (2011).

Qin, Xu; Xie, Wenchun; Su, Qi; Du, Wenzhe; Gross, Richard A. Protease-catalyzed oligomerization of L-lysine ethyl ester in aqueous solution, ACS Catalysis, 1(9), 1022-1034 (2011).

Liu, Jie; Jiang, Zhaozhong; Zhang, Shengmin; Liu, Chen; Gross, Richard A.; Kyriakides, Themis R.; Saltzman, W. Mark,  Biodegradation, biocompatibility, and drug delivery in poly(ω-pentadecalactone-co-p-dioxanone) copolyesters Biomaterials, 32(27), 6646-6654 (2011).

Kundu, Santanu; Bhangale, Atul S.; Wallace, William E.; Flynn, Kathleen M.; Guttman, Charles M.; Gross, Richard A.; Beers, Kathryn L. “Continuous Flow Enzyme-Catalyzed Polymerization in a Microreactor” Journal of the American Chemical Society 133(15), 6006-6011 (2011)

Yang, Yixin; Lu, Wenhua; Cai, Jiali; Hou, Yu; Ouyang, Suyang; Xie, Wenchun; Gross, Richard A. “Poly(oleic diacid-co-glycerol): comparison of polymer structure resulting from chemical and lipase catalysis” Macromolecules44(7), 1977-1985 (2011).

Lu, Wen-Hua; Ness, Jon E.; Xie, Wen-Chun; Zhang, Xiao-Yan; Minshull, Jeremy; Gross, Richard A.    “Biosynthesis of Monomers for Plastics from Renewable Oils” Journal of the American Chemical Society132(43), 15451-15455 (2010). 

Gualandi, Chiara; White, Lisa J.; Chen, Liu; Gross, Richard A.; Shakesheff, Kevin M.; Howdle, Steven M.; Scandola, Mariastella “Scaffold for tissue engineering fabricated by non-isothermal supercritical carbon dioxide foaming of a highly crystalline polyester” Acta Biomaterialia, 6(1), 130-136 (2010).

Focarete, Maria Letizia; Gualandi, Chiara; Scandola, Mariastella; Govoni, Marco; Giordano, Emanuele; Foroni, Laura; Valente, Sabrina; Pasquinelli, Gianandrea; Gao, Wei; Gross, Richard A. “Electrospun scaffolds of a polyhydroxyalkanoate consisting of ω-hydroxylpentadecanoate repeat units: fabrication and in vitro biocompatibility studies” Journal of Biomaterials Science, Polymer Edition21(10), 1283-1296 (2010). 

Viswanathan, Kodandaraman; Omorebokhae, Ruth; Li, Geng; Gross, Richard A. “Protease-Catalyzed Oligomerization of Hydrophobic Amino Acid Ethyl Esters in Homogeneous Reaction Media Using L-Phenylalanine as a Model System” Biomacromolecules, 11(8), 2152-2160 (2010). 

Gross, Richard A.; Ganesh, Manoj; Lu, Wenhua “Enzyme-catalysis breathes new life into polyester condensation polymerizations” Trends in Biotechnology, 28(8), 435-443 (2010). 

Viswanathan, Kodandaraman; Li, Geng; Gross, Richard A. “Protease catalyzed in situ C-terminal modification of oligoglutamate” Macromolecules43(12), 5245-5255(2010). 

Feder, David; Gross, Richard A. “Exploring Chain Length Selectivity in HiC-Catalyzed Polycondensation Reactions” Biomacromolecules11(3), 690-697 (2010). 

Liu, Z.; Gosser, Y.; Baker, P.J.; Ravee, Y.; Lu, Z.; Alemu, G.; Li, H.;  Butterfoss, G.L. Kong, X.-P. Gross, R.A.; Montclare, J.K. “Structural and Functional Studies of Aspergillus oryzae Cutinase: Enhanced Thermostability and Hydrolytic Activity of Synthetic Ester and Polyester Degradation” J. Am. Chem. Soc., 131 (43), pp 15711–15716 (2009)

Ganesh, M.; Dave, R.N.; L’Amoreaux, W.; Gross, R.A. “Embedded Enzymatic Biomaterial Degradation” Macromolecules, 42 (18), pp 6836–6839 (2009).

Ronkvist, A.M.; Lu, W.; Feder, D.; Gross, R.A. “Cutinase-Catalyzed Deacetylation of Poly(vinyl  acetate)” Macromolecules, 2009, 42 (16),  6086–6097 (2009)

Ronkvist, A.M.; Xie, W.; Lu, W.; Gross, R.A. “Cutinase-Catalyzed Hydrolysis of Poly(ethylene terephthalate)” Macromolecules, 2009, 42 (14), pp 5128–5138

Sleiman J.N., Kohlhoff S.A., Roblin P.A., Wallner S., Gross R., Hammerschlag M.R., Bluth M.H. “Sophorolipids as Antimicrobials” Ann. Clin. Lab Sci  39(1) 60-63.  

Chen, B., Pernodet, N., Rafailovich, M.H., Bakhtina, A. and Gross, R.A. Protein immobilization on epoxy-activated thin polymer films: Effect of surface wettability and enzyme loading Langmuir 24 (23): 13457-13464(2008).

Zini, E., Gazzano, M., Scandola, M., Wallner, S.R. and Gross, R. A. Glycolipid biomaterials: Solid-state properties of a poly(sophorolipid). Macromolecules 42(20): 7463-7468 (2008).

Li, G., Raman, V.K., Xie, W.C. and Gross R.A. Protease-catalyzed co-oligomerizations of L-leucine ethyl ester with L-glutamic acid diethyl ester: Sequence and chain length distributions. Macromolecules 41 (19): 7003-7012 (2008).

Fu, S.L., Wallner, S.R., Bowne, W.B., Hagler, M.D., Zenilman, M.E., Gross, R.A. and Bluth, M.H. Sophorolipids and their derivatives are lethal against human pancreatic cancer cells. Journal of Surgical Research 148 (1): 77-82 (2008).

Chen, B., Hu, J., Miller, E.M., Xie, W.H., Cai, M.M. and Gross, R.A. Candida antarctica lipase B chemically immobilized on epoxy-activated micro- and nanobeads: Catalysts for polyester synthesis. Biomacromolecules  9: 463-471(2008).

Dodds, D. R. and Gross, R. A. Chemicals from biomass. Science 318 (5854): 1250-1251 (2007).

Hardin, R., Pierre, J., Schulze, R., Mueller, C. M., Fu, S. L., Wallner, S. R., Stanek, A., Shah, V., Gross, R. A., Weedon, J., Nowakowski, M., Zenilman, M. E. and Bluth, M. H. Sophorolipids improve sepsis survival: Effects of dosing and derivatives. Journal of Surgical Research 142 (2): 314-319 (2007).

Fu, S. L., Mueller, C., Lin, Y. Y., Viterbo, D., Pierre, J., Shah, V., Gross, R., Schulze, R. and Zenilman, M. Sophorolipid treatment decreases LIPS induced inflammatory responses and NO production in macrophages. Journal of the American College of Surgeons 205 (3): S44-S44 (2007).

Kulshrestha, A. S., Gao, W., Fu, H. Y. and Gross, R. A. Synthesis and characterization of branched polymers from lipase-catalyzed trimethylolpropane copolymerizations. Biomacromolecules 8 (6): 1794-1801 (2007).

Chen, B., Miller, M. E. and Gross, R. A. Effects of porous polystyrene resin parameters on Candida antarctica Lipase B adsorption, distribution, and polyester synthesis activity. Langmuir  23 (11): 6467-6474 (2007).

Li, G., Vaidya, A., Viswanathan, K., Cui, J.R., Xie, W.C., Gao, W. and Gross, R.A. Rapid regioselective oligomerization of L-glutamic acid diethyl ester catalyzed by papain, Macromolecules 39 (23): 7915-7921 (2006)

Bluth, M.H., Kandil, E., Mueller, C.M., Shah, V., Lin, Y.Y., Zhang, H.,  Dresner, L., Lempert, L., Nowakowski, M., Gross, R.A., Schulze, R. and Zenilman, M.E.  Sophorolipids block lethal effects of septic shock in rats in a cecal ligation and puncture model of experimental sepsis. Critical Care Medicine 34 (1): 188-195 (2006)

Mueller, C.M., Viterbo, D., Murray, P.J., Shah, V., Gross, R.A., Schulze, R., Zenilman, M.E. and Bluth, M.H. Sophorolipid treatment decreases inflammatory cytokine expression in an in vitro model of experimental sepsis. Faseb Journal 20 (4): A204-A204 Part 1, (2006).

Shah V, Doncel GF, Seyoum T, Eaton KM, Zalenskaya I, Hagver R, Azim A, Gross R. Sophorolipids, microbial glycolipids with anti-human immunodeficiency virus and sperm-immobilizing activities. Antimicrob Agents Chemother; 149; 1-8 (2005).

Sahoo, B.; Brandstadt, K. F.; Lane, T. H.; Gross, R. A. “Sweet Silicones": Biocatalytic Reactions to Form Organosilicon Carbohydrate Macromers Org. Lett.; 7(18); 3857-3860 (2005).

Loos, K.; Kennedy, S. B.; Eidelman, N.; Tai, Y.; Zharnikov, M.; Amis, E. J.; Ulman, A.; Gross, R. A. Combinatorial Approach To Study Enzyme/Surface Interactions Langmuir; 21(12); 5237-5241 (2005).  

Kulshrestha, A. S.;  Gao, W.;  Gross, R.A. “Glycerol Copolyesters: Control of Branching and Molecular Weight Using a Lipase Catalyst”, Macromolecules, (2005); 38(8); 3193-3204

Zhang, L., Somasundaran, P., Singh, S. K.,  Felse, A. P., Gross, R.A. Synthesis and interfacial properties of sophorolipid derivatives Colloids and Surfaces A: Physicochem.Eng. Aspects; 240; 75-82 (2004)

Zhou, S., Xu, Chang.., Wang, J., Gao, W., Akhverdiyeva.,  Shah, V., Gross,  R. A. Supramolecular Assembles of a Naturally Derived Sopholipid. Langmuir; 20; 7926-7932 (2004)

Mei, Y., Kumar, A, Gao, W, Gross, R.A., Kennedy, S.B., Washburn, N.R., Amis, E,.J., Elliot, John T.  Biocompatibility of sorbitol-containing polyesters. Part 1: Synthesis, surface analysis and cell response in vitro; Biomaterials; 25; 4195-4201 (2004)

Singh, S,K,  Felse, A. P., Nunez, A., Foglia, T.A. and  Gross, R.A. Regioselective Enzyme-Catalyzed Synthesis of Sophorolipid Esters, Amides and Multifunctional Monomers. J. Org. Chem.; 68; 5466-5477 (2003)

Kumar, A.; Kulshrestha, A. S.; Gao, W.; Gross, R. A.; Versatile Route to Polyol Polyesters by Lipase Catalysis Macromolecules; 36(22); 8219-8221 (2003). 

Mei, Y.; Miller, L.; Gao, W.; Gross, R. A.; Imaging the Distribution and Secondary Structure of Immobilized Enzymes Using Infrared Microspectroscopy Biomacromolecules; 4(1); 70-74 (2003).

Dyal, A., Loos, Katja., Noto, M., Chang, S.W., Spagnoli, C., Shafi, Kurikka V.P.M., Ulman, A., Cowman,M.,Gross. R.A. Activity of Candida rugosa Lipase Immobilized on g-Fe2O3 Magnetic Nanoparticles  J. Am. Chem. Soc.; 125; 1684-1685 (2003).

Mei, Y, Kumar, A, Gross; R.A., “Kinetics and Mechanism of Candida antarctica Lipase B Catalyzed Solution Polymerization of ε-Caprolactone”, Macromolecules; 36(15); 5530-5536 (2003).

Gross, R. A., Kalra, B; “Biodegradable Polymers for the Environment”, Science, 297, 803-806 (2002).

R. Kumar and R.A. Gross, “Biocatalytic Route to Well-Defined Macromers Built around a Sugar Core, J. Am. Chem. Soc., 124(9) (2002).   

V Guilmanov, A Ballistreri, G Impallomeni, R.A. Gross, “Oxygen Transfer Rate and Sophorose Lipid Production by Candida bombicola”, Biotechnol. and Bioeng; 77(5), 489-494 (2002)

R. A. Gross, A Kumar, B Kalra, "In-vitro Enzyme Catalyzed Polymer Synthesis", Chemical Reviews, 101(7), 2097-2124 (2001).

J.W. Lee, F. Dang, W.G. Yeomans, A.L. Allen, R.A. Gross, D.L. Kaplan, “Acetobacter xylinium ATCC 10245: Production of Chitosan-Cellulose and Chitin-Cellulose Exopolymers, Applied and Environmental Microbiology, 67(9), 3970-3975 (2001).

A Kumar, R.A Gross, “Lipase-Catalyzed Transesterification: New Synthetic Routes To Copolyesters”, J. Am. Chem. Soc.; 122; 11767-11770 (2000).
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