Christian Isak Jørgensen

Linking Phospholipase Mobility to Activity by Single-Molecule Wide-Field Microscopy

Many of the biological processes taking place in cells are mediated by enzymatic reactions occurring in the cell membrane. Understanding interfacial enzymatic catalysis is therefore crucial to the understanding of cellular function. Unfortunately, a full picture of the overall mechanism of interfacial enzymatic catalysis, and particularly the important diffusion processes therein, remains unresolved. Herein we demonstrate that single-molecule wide-field fluorescence microscopy can yield important new information on these processes. We image phospholipase enzymes acting upon bilayers of their natural phospholipid substrate, tracking the diffusion of thousands of individual enzymes while simultaneously visualising local structural changes to the substrate layer. We study several enzyme types with different affinities and catalytic activities towards the substrate. Analysis of the trajectories of each enzyme type allows us successfully to correlate the mobility of phospholipase with its catalytic activity at the substrate. The methods introduced herein represent a promising new approach to the study of interfacial/heterogeneous catalysis systems.

Linking hydrolysis performance to Trichoderma reesei cellulolytic enzyme profile

Trichoderma reesei expresses a large number of enzymes involved in lignocellulose hydrolysis and the mechanism of how these enzymes work together is too complex to study by traditional methods, for example, by spiking with single enzymes and monitoring hydrolysis performance. In this study, a multivariate approach, partial least squares regression, was used to see whether it could help explain the correlation between enzyme profile and hydrolysis performance. Diverse enzyme mixtures were produced by T. reesei Rut‐C30 by exploiting various fermentation conditions and used for hydrolysis of washed pretreated corn stover as a measure of enzyme performance. In addition, the enzyme mixtures were analyzed by liquid chromatography–tandem mass spectrometry to identify and quantify the different proteins. A multivariate model was applied for the prediction of enzyme performance based on the combination of different proteins present in an enzyme mixture. The multivariate model was used for identification of candidate proteins that are correlated to enzyme performance on pretreated corn stover. A very large variation in hydrolysis performance was observed and this was clearly caused by the difference in fermentation conditions. Besides β‐glucosidase, the multivariate model identified several xylanases, Cip1 and Cip2, as relevant proteins to study further. Biotechnol. Bioeng. 2016;113: 1001–1010.

Phosphate Selective Uranyl Photo‐Affinity Cleavage of Proteins. Determination of Phosphorylation Sites

Phosphorylation of proteins is one of the most important mechanisms in cellular signaling and is involved in cellular processes such as metabolism, transcription, translation, cell cycle, movement, apoptosis, and differentiation. Phosphorylation takes place at serine, threonine, and tyrosine residues in a 1000:100:1 ratio, and it has been estimated that 30% of all proteins are reversibly phosphorylated at one or multiple sites at some point during their lifetime. This is facilitated by the high number of kinases and phosphatases, which constitute 2% of all human genome genes. Phosphorylation of proteins is a reversible process, and proteins can be phosphorylated at substoichiometric levels. For proteins containing multiple phosphorylation sites, each site can be associated with a different function; this makes the particular function in question ACHTUNGTRENNUNGdependent on the phosphorylation pattern. To understand the role of dynamic protein phosphorylation, the identification of the exact sites and extent of phosphorylation is crucial. At present, state-of-the-art techniques for investigation of protein phosphorylation make use of mass spectrometry (MS), subsequent to protease digests (most often trypsin) of an isolated phosphoprotein or of the entire phosphoproteome in a cell lysate. Secondly, phosphopeptide enrichment is carried out to reduce the number of peptides to be analyzed. Finally N-terminal sequencing and/or MS analysis are performed to identify the specific positions of phosphorylation. Although different MS approaches have been used for detecting phosphorylated positions in the proteome, limitations and difficulties remain concerning signal suppression of phosphate containing peptides, dephosphorylation, difficulties in achieving coverage of the full length of long peptides, peptides present in low amounts, peptides phosphorylated at substoichiometric levels, and finally, difficulties in distinguishing among multiple possible phosphorylation sites within in a given peptide fragment. Thus, analysis of peptides after trypsin digests is not straightforward and determination of the exact site of phosphorylation often fails for recovered phosphopepACHTUNGTRENNUNGtides. Therefore alternative approaches have been considered in order to develop improved methods for phosphorylation site determination in the proteome. An attractive goal has been to develop phosphospecific proteases in order to reduce the amount of peptide products, which have to be analyzed. Furthermore, such a protease would generate peptide products containing the phosphoamino acid residue positioned either at the N or C terminus thereby significantly simplifying MS analysis and the identification of the phosphorylation sites. Although no phosphospecific protease has so far been found in nature, a chemical approach has been developed where phosphoserines and phosphothreonines are converted into lysine analogues (amino-ethyl cysteine and b-methyl S-ethyl cysteine) to generate cleavage sites for lysine specific proteases. However, this methodology is both complex and laborious and involves several chemical and enzymatic modifications, and most importantly, induces problems in distinguishing naturally occurring lysines from those generated from phosphorylated serine and threonine residues. Uranyl photocleavage has for two decades been used for studying protein–double-stranded (ds) DNA interactions, drug– dsDNA interactions, and the interactions of metal-ions with nucleic acids. Uranyl photocleavage of nucleic acids is based on the high affinity of the divalent uranyl cation for phosphates in the backbone of nucleic acids and the strong oxidation power of the excited state of the uranyl ion. Upon excitation of the bound uranyl ion, the high oxidation potential of uranyl induces breakage at the sugar-phosphate backbone at each nucleotide. It was recently reported that proteins can also be photocleaved by uranyl but at low efficiency. In view of the high affinity of the uranyl(VI) ion for phosphate, we have now systematically analyzed whether phosphorylated sites present in proteins could recruit the uranyl ion for subsequent cleavage. We find that both the specificity as well as the efficiency of the photocleavage reaction is increased in the presence of phosphorylated residues and this opens the potential application for detection of phosphorylation sites in proteins by uranyl photocleavage. To examine whether site-specific cleavage at phoshorylated residues in proteins could be induced by uranyl photocleavage, three different phosphoproteins were selected as model systems: a-casein, b-casein, and ovalbumin. The three proteins differ in size, structure, and extent of phosphorylation, and represent different patterns of phosphorylation in proteins. Initially all three proteins were subjected to uranyl photocleavage at different uranyl/protein ratios and irradiated at 320 nm. The [a] L. H. Kristensen, Prof. P. E. Nielsen, N. E. Møllegaard Department of Cellular and Molecular Medicine Panum Institute, University of Copenhagen Blegdamsvej 3, 2200 Copenhagen N (Denmark) Fax: (+45)35-32-77-32 E-mail : nem@imbg.ku.dk [b] C. I. Jørgensen Novozymes A/S Novo Alle 6B 3.90, 2880 Bagsværd (Denmark) [c] B. B. Kragelund Department of Biology, Structural Biology and NMR Laboratory University of Copenhagen Ole Maaløes Vej 5, 2200 Copenhagen N (Denmark) Supporting information for this article is available on the WWW under http://www.chembiochem.org or from the author.