In this study, we describe an effective protocol for use in a multiplexed high-throughput antibody microarray with glycan binding protein detection that allows for the glycosylation profiling of specific proteins. Glycosylation of proteins is the most prevalent post-translational modification found on proteins, and leads diversified modifications of the physical, chemical, and biological properties of proteins. Because the glycosylation machinery is particularly susceptible to disease progression and malignant transformation, aberrant glycosylation has been recognized as early detection biomarkers for cancer and other diseases. However, current methods to study protein glycosylation typically are too complicated or expensive for use in most normal laboratory or clinical settings and a more practical method to study protein glycosylation is needed. The new protocol described in this study makes use of a chemically blocked antibody microarray with glycan-binding protein (GBP) detection and significantly reduces the time, cost, and lab equipment requirements needed to study protein glycosylation. In this method, multiple immobilized glycoprotein-specific antibodies are printed directly onto the microarray slides and the N-glycans on the antibodies are blocked. The blocked, immobilized glycoprotein-specific antibodies are able to capture and isolate glycoproteins from a complex sample that is applied directly onto the microarray slides. Glycan detection then can be performed by the application of biotinylated lectins and other GBPs to the microarray slide, while binding levels can be determined using Dylight 549-Streptavidin. Through the use of an antibody panel and probing with multiple biotinylated lectins, this method allows for an effective glycosylation profile of the different proteins found in a given human or animal sample to be developed.
Glycosylation of protein, which is the most ubiquitous post-translational modification on proteins, modifies the physical, chemical, and biological properties of a protein, and plays a fundamental role in various biological processes1-6. Because the glycosylation machinery is particularly susceptible to disease progression and malignant transformation, aberrant glycosylation has been recognized as early detection biomarkers for cancer and other diseases 7-12. In fact, most current cancer biomarkers, such as the L3 fraction of α-1 fetoprotein (AFP) for hepatocellular carcinoma 13-15, and CA199 for pancreatic cancer 16, 17 are all aberrant glycan moieties on glycoproteins. However, methods to study protein glycosylation have been complicated, and not suitable for routine laboratory and clinical settings. Chen et al. has recently invented a chemically blocked antibody microarray with a glycan-binding protein (GBP) detection method for high-throughput and multiplexed profile glycosylation of native glycoproteins in a complex sample 18. In this affinity based microarray method, multiple immobilized glycoprotein-specific antibodies capture and isolate glycoproteins from the complex mixture directly on the microarray slide, and the glycans on each individual captured protein are measured by GBPs. Because all normal antibodies contain N-glycans which could be recognized by most GBPs, the critical step of this method is to chemically block the glycans on the antibodies from binding to GBP. In the procedure, the cis-diol groups of the glycans on the antibodies were first oxidized to aldehyde groups by using NaIO4 in sodium acetate buffer avoiding light. The aldehyde groups were then conjugated to the hydrazide group of a cross-linker, 4-(4-N-MaleimidoPhenyl)butyric acid Hydrazide HCl (MPBH), followed by the conjugation of a dipeptide, Cys-Gly, to the maleimide group of the MPBH. Thus, the cis-diol groups on glycans of antibodies were converted into bulky none hydroxyl groups, which hindered the lectins and other GBPs bindings to the capture antibodies. This blocking procedure makes the GBPs and lectins bind only to the glycans of captured proteins. After this chemically blocking, serum samples were incubated with the antibody microarray, followed by the glycans detection by using different biotinylated lectins and GBPs, and visualized with Cy3-streptavidin. The parallel use of an antibody panel and multiple lectin probing provides discrete glycosylation profiles of multiple proteins in a given sample 18-20. This method has been used successfully in multiple different labs 1, 7, 13, 19-31. However, stability of MPBH and Cys-Gly, complicated and extended procedure in this method affect the reproducibility, effectiveness and efficiency of the method. In this new protocol, we replaced both MPBH and Cys-Gly with one much more stable reagent glutamic acid hydrazide (Glu-hydrazide), which significantly improved the reproducibility of the method, simplified and shorten the whole procedure so that the it can be completed within one working day. In this new protocol, we describe the detailed procedure of the protocol which can be readily adopted by normal labs for routine protein glycosylation study and techniques which are necessary to obtain reproducible and repeatable results.
21 Related JoVE Articles!
Glycopeptide Capture for Cell Surface Proteomics
Institutions: Simon Fraser University.
Cell surface proteins, including extracellular matrix proteins, participate in all major cellular processes and functions, such as growth, differentiation, and proliferation. A comprehensive characterization of these proteins provides rich information for biomarker discovery, cell-type identification, and drug-target selection, as well as helping to advance our understanding of cellular biology and physiology. Surface proteins, however, pose significant analytical challenges, because of their inherently low abundance, high hydrophobicity, and heavy post-translational modifications. Taking advantage of the prevalent glycosylation on surface proteins, we introduce here a high-throughput glycopeptide-capture approach that integrates the advantages of several existing N-glycoproteomics means. Our method can enrich the glycopeptides derived from surface proteins and remove their glycans for facile proteomics using LC-MS. The resolved N-glycoproteome comprises the information of protein identity and quantity as well as their sites of glycosylation. This method has been applied to a series of studies in areas including cancer, stem cells, and drug toxicity. The limitation of the method lies in the low abundance of surface membrane proteins, such that a relatively large quantity of samples is required for this analysis compared to studies centered on cytosolic proteins.
Molecular Biology, Issue 87, membrane protein, N-linked glycoprotein, post-translational modification, mass spectrometry, HPLC, hydrazide chemistry, N-glycoproteomics, glycopeptide capture
Lectin-based Isolation and Culture of Mouse Embryonic Motoneurons
Institutions: Ruhr-University Bochum, University of Wuerzburg.
Spinal motoneurons develop towards postmitotic stages through early embryonic nervous system development and subsequently grow out dendrites and axons. Neuroepithelial cells of the neural tube that express Nkx6.1 are the unique precursor cells for spinal motoneurons1
. Though postmitotic motoneurons move towards their final position and organize themselves into columns along the spinal tract2,3
. More than 90% of all these differentiated and positioned motoneurons express the transcription factors Islet 1/2. They innervate the muscles of the limbs as well as those of the body and the inner organs. Among others, motoneurons typically express the high affinity receptors for brain derived neurotrophic factor (BDNF) and Neurotrophin-3 (NT-3), the tropomyosin-related kinase B and C (TrkB, TrkC). They do not express the tropomyosin-related kinase A (TrkA)4
. Beside the two high affinity receptors, motoneurons do express the low affinity neurotrophin receptor p75NTR
. The p75NTR
can bind all neurotrophins with similar but lower affinity to all neurotrophins than the high affinity receptors would bind the mature neurotrophins. Within the embryonic spinal cord, the p75NTR
is exclusively expressed by the spinal motoneurons5
. This has been used to develop motoneuron isolation techniques to purify the cells from the vast majority of surrounding cells6
. Isolating motoneurons with the help of specific antibodies (panning) against the extracellular domains of p75NTR
has turned out to be an expensive method as the amount of antibody used for a single experiment is high due to the size of the plate used for panning. A much more economical alternative is the use of lectin. Lectin has been shown to specifically bind to p75NTR
. The following method describes an alternative technique using wheat germ agglutinin for a preplating procedure instead of the p75NTR
antibody. The lectin is an extremely inexpensive alternative to the p75NTR
antibody and the purification grades using lectin are comparable to that of the p75NTR
antibody. Motoneurons from the embryonic spinal cord can be isolated by this method, survive and grow out neurites.
Neuroscience, Issue 55, p75NTR, spinal cord, lectin, axon, dendrite
Pulse-chase Analysis of N-linked Sugar Chains from Glycoproteins in Mammalian Cells
Institutions: Tel Aviv University.
Attachment of the Glc3
precursor oligosaccharide to nascent polypeptides in the ER is a common modification for secretory proteins. Although this modification was implicated in several biological processes, additional aspects of its function are emerging, with recent evidence of its role in the production of signals for glycoprotein quality control and trafficking. Thus, phenomena related to N-linked glycans and their processing are being intensively investigated. Methods that have been recently developed for proteomic analysis have greatly improved the characterization of glycoprotein N-linked glycans. Nevertheless, they do not provide insight into the dynamics of the sugar chain processing involved. For this, labeling and pulse-chase analysis protocols are used that are usually complex and give very low yields. We describe here a simple method for the isolation and analysis of metabolically labeled N-linked oligosaccharides. The protocol is based on labeling of cells with [2-3
H] mannose, denaturing lysis and enzymatic release of the oligosaccharides from either a specifically immunoprecipitated protein of interest or from the general glycoprotein pool by sequential treatments with endo H and N-glycosidase F, followed by molecular filtration (Amicon). In this method the isolated oligosaccharides serve as an input for HPLC analysis, which allows discrimination between various glycan structures according to the number of monosaccharide units comprising them, with a resolution of a single monosaccharide. Using this method we were able to study high mannose N-linked oligosaccharide profiles of total cell glycoproteins after pulse-chase in normal conditions and under proteasome inhibition. These profiles were compared to those obtained from an immunoprecipitated ER-associated degradation (ERAD) substrate. Our results suggest that most NIH 3T3 cellular glycoproteins are relatively stable and that most of their oligosaccharides are trimmed to Man9-8
. In contrast, unstable ERAD substrates are trimmed to Man6-5
and glycoproteins bearing these species accumulate upon inhibition of proteasomal degradation.
Cellular Biology, Issue 38, N-linked oligosaccharide, mannose-labeling, endoplasmic reticulum associated degradation, calnexin, glycosylation, mannosidase
Reconstitution Of β-catenin Degradation In Xenopus Egg Extract
Institutions: Vanderbilt University Medical Center, Cincinnati Children's Hospital Medical Center, Vanderbilt University School of Medicine.
egg extract is a well-characterized, robust system for studying the biochemistry of diverse cellular processes. Xenopus
egg extract has been used to study protein turnover in many cellular contexts, including the cell cycle and signal transduction pathways1-3
. Herein, a method is described for isolating Xenopus
egg extract that has been optimized to promote the degradation of the critical Wnt pathway component, β-catenin. Two different methods are described to assess β-catenin protein degradation in Xenopus
egg extract. One method is visually informative ([35
S]-radiolabeled proteins), while the other is more readily scaled for high-throughput assays (firefly luciferase-tagged fusion proteins). The techniques described can be used to, but are not limited to, assess β-catenin protein turnover and identify molecular components contributing to its turnover. Additionally, the ability to purify large volumes of homogenous Xenopus
egg extract combined with the quantitative and facile readout of luciferase-tagged proteins allows this system to be easily adapted for high-throughput screening for modulators of β-catenin degradation.
Molecular Biology, Issue 88, Xenopus laevis, Xenopus egg extracts, protein degradation, radiolabel, luciferase, autoradiography, high-throughput screening
Blastomere Explants to Test for Cell Fate Commitment During Embryonic Development
Institutions: The George Washington University, The George Washington University.
Fate maps, constructed from lineage tracing all of the cells of an embryo, reveal which tissues descend from each cell of the embryo. Although fate maps are very useful for identifying the precursors of an organ and for elucidating the developmental path by which the descendant cells populate that organ in the normal embryo, they do not illustrate the full developmental potential of a precursor cell or identify the mechanisms by which its fate is determined. To test for cell fate commitment, one compares a cell's normal repertoire of descendants in the intact embryo (the fate map) with those expressed after an experimental manipulation. Is the cell's fate fixed (committed) regardless of the surrounding cellular environment, or is it influenced by external factors provided by its neighbors? Using the comprehensive fate maps of the Xenopus
embryo, we describe how to identify, isolate and culture single cleavage stage precursors, called blastomeres. This approach allows one to assess whether these early cells are committed to the fate they acquire in their normal environment in the intact embryo, require interactions with their neighboring cells, or can be influenced to express alternate fates if exposed to other types of signals.
Developmental Biology, Issue 71, Cellular Biology, Molecular Biology, Anatomy, Physiology, Biochemistry, Xenopus laevis, fate mapping, lineage tracing, cell-cell signaling, cell fate, blastomere, embryo, in situ hybridization, animal model
Fluorescence Microscopy Methods for Determining the Viability of Bacteria in Association with Mammalian Cells
Institutions: University of Virginia Health Sciences Center.
Central to the field of bacterial pathogenesis is the ability to define if and how microbes survive after exposure to eukaryotic cells. Current protocols to address these questions include colony count assays, gentamicin protection assays, and electron microscopy. Colony count and gentamicin protection assays only assess the viability of the entire bacterial population and are unable to determine individual bacterial viability. Electron microscopy can be used to determine the viability of individual bacteria and provide information regarding their localization in host cells. However, bacteria often display a range of electron densities, making assessment of viability difficult. This article outlines protocols for the use of fluorescent dyes that reveal the viability of individual bacteria inside and associated with host cells. These assays were developed originally to assess survival of Neisseria gonorrhoeae
in primary human neutrophils, but should be applicable to any bacterium-host cell interaction. These protocols combine membrane-permeable fluorescent dyes (SYTO9 and 4',6-diamidino-2-phenylindole [DAPI]), which stain all bacteria, with membrane-impermeable fluorescent dyes (propidium iodide and SYTOX Green), which are only accessible to nonviable bacteria. Prior to eukaryotic cell permeabilization, an antibody or fluorescent reagent is added to identify extracellular bacteria. Thus these assays discriminate the viability of bacteria adherent to and inside eukaryotic cells. A protocol is also provided for using the viability dyes in combination with fluorescent antibodies to eukaryotic cell markers, in order to determine the subcellular localization of individual bacteria. The bacterial viability dyes discussed in this article are a sensitive complement and/or alternative to traditional microbiology techniques to evaluate the viability of individual bacteria and provide information regarding where bacteria survive in host cells.
Microbiology, Issue 79, Immunology, Infection, Cancer Biology, Genetics, Cellular Biology, Molecular Biology, Medicine, Biomedical Engineering, Microscopy, Confocal, Microscopy, Fluorescence, Bacteria, Bacterial Infections and Mycoses, bacteria, infection, viability, fluorescence microscopy, cell, imaging
Two Types of Assays for Detecting Frog Sperm Chemoattraction
Institutions: University of Illinois, Urbana-Champaign, Arizona State University .
Sperm chemoattraction in invertebrates can be sufficiently robust that one can place a pipette containing the attractive peptide into a sperm suspension and microscopically visualize sperm accumulation around the pipette1
. Sperm chemoattraction in vertebrates such as frogs, rodents and humans is more difficult to detect and requires quantitative assays. Such assays are of two major types - assays that quantitate sperm movement to a source of chemoattractant, so-called sperm accumulation assays, and assays that actually track the swimming trajectories of individual sperm.
Sperm accumulation assays are relatively rapid allowing tens or hundreds of assays to be done in a single day, thereby allowing dose response curves and time courses to be carried out relatively rapidly. These types of assays have been used extensively to characterize many well established chemoattraction systems - for example, neutrophil chemotaxis to bacterial peptides and sperm chemotaxis to follicular fluid. Sperm tracking assays can be more labor intensive but offer additional data on how chemoattractancts actually alter the swimming paths that sperm take. This type of assay is needed to demonstrate the orientation of sperm movement relative to the chemoattrractant gradient axis and to visualize characteristic turns or changes in orientation that bring the sperm closer to the egg.
Here we describe methods used for each of these two types of assays. The sperm accumulation assay utilized is called a "two-chamber" assay. Amphibian sperm are placed in a tissue culture plate insert with a polycarbonate filter floor having 12 μm diameter pores. Inserts with sperm are placed into tissue culture plate wells containing buffer and a chemoatttractant carefully pipetted into the bottom well where the floor meets the wall (see Fig. 1). After incubation, the top insert containing the sperm reservoir is carefully removed, and sperm in the bottom chamber that have passed through the membrane are removed, pelleted and then counted by hemocytometer or flow cytometer.
The sperm tracking assay utilizes a Zigmond chamber originally developed for observing neutrophil chemotaxis and modified for observation of sperm by Giojalas and coworkers2,3
. The chamber consists of a thick glass slide into which two vertical troughs have been machined. These are separated by a 1 mm wide observation platform. After application of a cover glass, sperm are loaded into one trough, the chemoattractant agent into the other and movement of individual sperm visualized by video microscopy. Video footage is then analyzed using software to identify two-dimensional cell movements in the x-y plane as a function of time (xyt data sets) that form the trajectory of each sperm.
Developmental Biology, Issue 58, Sperm chemotaxis, fertilization, sperm accumulation assay, sperm tracking assay, sperm motility, Xenopus laevis, egg jelly
Measuring Cation Transport by Na,K- and H,K-ATPase in Xenopus Oocytes by Atomic Absorption Spectrophotometry: An Alternative to Radioisotope Assays
Institutions: Technical University of Berlin, Oregon Health & Science University.
Whereas cation transport by the electrogenic membrane transporter Na+
-ATPase can be measured by electrophysiology, the electroneutrally operating gastric H+
-ATPase is more difficult to investigate. Many transport assays utilize radioisotopes to achieve a sufficient signal-to-noise ratio, however, the necessary security measures impose severe restrictions regarding human exposure or assay design. Furthermore, ion transport across cell membranes is critically influenced by the membrane potential, which is not straightforwardly controlled in cell culture or in proteoliposome preparations. Here, we make use of the outstanding sensitivity of atomic absorption spectrophotometry (AAS) towards trace amounts of chemical elements to measure Rb+
transport by Na+
- or gastric H+
-ATPase in single cells. Using Xenopus
oocytes as expression system, we determine the amount of Rb+
) transported into the cells by measuring samples of single-oocyte homogenates in an AAS device equipped with a transversely heated graphite atomizer (THGA) furnace, which is loaded from an autosampler. Since the background of unspecific Rb+
uptake into control oocytes or during application of ATPase-specific inhibitors is very small, it is possible to implement complex kinetic assay schemes involving a large number of experimental conditions simultaneously, or to compare the transport capacity and kinetics of site-specifically mutated transporters with high precision. Furthermore, since cation uptake is determined on single cells, the flux experiments can be carried out in combination with two-electrode voltage-clamping (TEVC) to achieve accurate control of the membrane potential and current. This allowed e.g.
to quantitatively determine the 3Na+
transport stoichiometry of the Na+
-ATPase and enabled for the first time to investigate the voltage dependence of cation transport by the electroneutrally operating gastric H+
-ATPase. In principle, the assay is not limited to K+
-transporting membrane proteins, but it may work equally well to address the activity of heavy or transition metal transporters, or uptake of chemical elements by endocytotic processes.
Biochemistry, Issue 72, Chemistry, Biophysics, Bioengineering, Physiology, Molecular Biology, electrochemical processes, physical chemistry, spectrophotometry (application), spectroscopic chemical analysis (application), life sciences, temperature effects (biological, animal and plant), Life Sciences (General), Na+,K+-ATPase, H+,K+-ATPase, Cation Uptake, P-type ATPases, Atomic Absorption Spectrophotometry (AAS), Two-Electrode Voltage-Clamp, Xenopus Oocytes, Rb+ Flux, Transversely Heated Graphite Atomizer (THGA) Furnace, electrophysiology, animal model
Identification and Characterization of Protein Glycosylation using Specific Endo- and Exoglycosidases
Institutions: New England Biolabs.
Glycosylation, the addition of covalently linked sugars, is a major post-translational modification of proteins that can significantly affect processes such as cell adhesion, molecular trafficking, clearance, and signal transduction1-4
. In eukaryotes, the most common glycosylation modifications in the secretory pathway are additions at consensus asparagine residues (N
-linked); or at serine or threonine residues (O
-linked) (Figure 1). Initiation of N
-glycan synthesis is highly conserved in eukaryotes, while the end products can vary greatly among different species, tissues, or proteins. Some glycans remain unmodified ("high mannose N
-glycans") or are further processed in the Golgi ("complex N
-glycans"). Greater diversity is found for O
-glycans, which start with a common N
-Acetylgalactosamine (GalNAc) residue in animal cells but differ in lower organisms1
The detailed analysis of the glycosylation of proteins is a field unto itself and requires extensive resources and expertise to execute properly. However a variety of available enzymes that remove sugars (glycosidases) makes possible to have a general idea of the glycosylation status of a protein in a standard laboratory setting. Here we illustrate the use of glycosidases for the analysis of a model glycoprotein: recombinant human chorionic gonadotropin beta (hCGβ), which carries two N
-glycans and four O
. The technique requires only simple instrumentation and typical consumables, and it can be readily adapted to the analysis of multiple glycoprotein samples.
Several enzymes can be used in parallel to study a glycoprotein. PNGase F is able to remove almost all types of N
. For O
-glycans, there is no available enzyme that can cleave an intact oligosaccharide from the protein backbone. Instead, O
-glycans are trimmed by exoglycosidases to a short core, which is then easily removed by O
-Glycosidase. The Protein Deglycosylation Mix contains PNGase F, O
-Glycosidase, Neuraminidase (sialidase), β1-4 Galactosidase, and β-N
-Acetylglucosaminidase. It is used to simultaneously remove N
-glycans and some O
. Finally, the Deglycosylation Mix was supplemented with a mixture of other exoglycosidases (α-N
-Acetylgalactosaminidase, α1-2 Fucosidase, α1-3,6 Galactosidase, and β1-3 Galactosidase ), which help remove otherwise resistant monosaccharides that could be present in certain O
SDS-PAGE/Coomasie blue is used to visualize differences in protein migration before and after glycosidase treatment. In addition, a sugar-specific staining method, ProQ Emerald-300, shows diminished signal as glycans are successively removed. This protocol is designed for the analysis of small amounts of glycoprotein (0.5 to 2 μg), although enzymatic deglycosylation can be scaled up to accommodate larger quantities of protein as needed.
Molecular Biology , Issue 58, Glycoprotein, N-glycan, O-glycan, PNGase F, O-glycosidase, deglycosylation, glycosidase
Ablation of a Single Cell From Eight-cell Embryos of the Amphipod Crustacean Parhyale hawaiensis
Institutions: Harvard University.
The amphipod Parhyale hawaiensis
is a small crustacean found in intertidal marine habitats worldwide. Over the past decade, Parhyale
has emerged as a promising model organism for laboratory studies of development, providing a useful outgroup comparison to the well studied arthropod model organism Drosophila melanogaster
. In contrast to the syncytial cleavages of Drosophila
, the early cleavages of Parhyale
are holoblastic. Fate mapping using tracer dyes injected into early blastomeres have shown that all three germ layers and the germ line are established by the eight-cell stage. At this stage, three blastomeres are fated to give rise to the ectoderm, three are fated to give rise to the mesoderm, and the remaining two blastomeres are the precursors of the endoderm and germ line respectively. However, blastomere ablation experiments have shown that Parhyale
embryos also possess significant regulatory capabilities, such that the fates of blastomeres ablated at the eight-cell stage can be taken over by the descendants of some of the remaining blastomeres. Blastomere ablation has previously been described by one of two methods: injection and subsequent activation of phototoxic dyes or manual ablation. However, photoablation kills blastomeres but does not remove the dead cell body from the embryo. Complete physical removal of specific blastomeres may therefore be a preferred method of ablation for some applications. Here we present a protocol for manual removal of single blastomeres from the eight-cell stage of Parhyale
embryos, illustrating the instruments and manual procedures necessary for complete removal of the cell body while keeping the remaining blastomeres alive and intact. This protocol can be applied to any Parhyale
cell at the eight-cell stage, or to blastomeres of other early cleavage stages. In addition, in principle this protocol could be applicable to early cleavage stage embryos of other holoblastically cleaving marine invertebrates.
Developmental Biology, Issue 85, Amphipod, experimental embryology, micromere, germ line, ablation, developmental potential, vasa
Glycan Profiling of Plant Cell Wall Polymers using Microarrays
Institutions: University of Melbourne, University of Melbourne, CSIRO Plant Industry, Black Mountain Laboratories, University of Copenhagen.
Plant cell walls are complex matrixes of heterogeneous glycans which play an important role in the physiology and development of plants and provide the raw materials for human societies (e.g.
wood, paper, textile and biofuel industries)1,2
. However, understanding the biosynthesis and function of these components remains challenging.
Cell wall glycans are chemically and conformationally diverse due to the complexity of their building blocks, the glycosyl residues. These form linkages at multiple positions and differ in ring structure, isomeric or anomeric configuration, and in addition, are substituted with an array of non-sugar residues. Glycan composition varies in different cell and/or tissue types or even sub-domains of a single cell wall3
. Furthermore, their composition is also modified during development1
, or in response to environmental cues4
In excess of 2,000 genes have Plant cell walls are complex matrixes of heterogeneous glycans been predicted to be involved in cell wall glycan biosynthesis and modification in Arabidopsis5
. However, relatively few of the biosynthetic genes have been functionally characterized 4,5
. Reverse genetics approaches are difficult because the genes are often differentially expressed, often at low levels, between cell types6
. Also, mutant studies are often hindered by gene redundancy or compensatory mechanisms to ensure appropriate cell wall function is maintained7
. Thus novel approaches are needed to rapidly characterise the diverse range of glycan structures and to facilitate functional genomics approaches to understanding cell wall biosynthesis and modification.
Monoclonal antibodies (mAbs)8,9
have emerged as an important tool for determining glycan structure and distribution in plants. These recognise distinct epitopes present within major classes of plant cell wall glycans, including pectins, xyloglucans, xylans, mannans, glucans and arabinogalactans. Recently their use has been extended to large-scale screening experiments to determine the relative abundance of glycans in a broad range of plant and tissue types simultaneously9,10,11
Here we present a microarray-based glycan screening method called Comprehensive Microarray Polymer Profiling (CoMPP) (Figures 1 & 2
that enables multiple samples (100 sec) to be screened using a miniaturised microarray platform with reduced reagent and sample volumes. The spot signals on the microarray can be formally quantified to give semi-quantitative data about glycan epitope occurrence. This approach is well suited to tracking glycan changes in complex biological systems12
and providing a global overview of cell wall composition particularly when prior knowledge of this is unavailable.
Plant Biology, Issue 70, Molecular Biology, Cellular Biology, Genetics, Genomics, Proteomics, Proteins, Cell Walls, Polysaccharides, Monoclonal Antibodies, Microarrays, CoMPP, glycans, Arabidopsis, tissue collection
A Lectin HPLC Method to Enrich Selectively-glycosylated Peptides from Complex Biological Samples
Institutions: University of California, San Francisco - UCSF, Buck Institute for Age Research, Purdue University.
Glycans are an important class of post-translational modifications. Typically found on secreted and extracellular molecules, glycan structures signal the internal status of the cell. Glycans on tumor cells tend to have abundant sialic acid and fucose moieties. We propose that these cancer-associated glycan variants be exploited for biomarker development aimed at diagnosing early-stage disease. Accordingly, we developed a mass spectrometry-based workflow that incorporates chromatography on affinity matrices formed from lectins, proteins that bind specific glycan structures. The lectins Sambucus nigra (SNA) and Aleuria aurantia (AAL), which bind sialic acid and fucose, respectively, were covalently coupled to POROS beads (Applied Biosystems) and packed into PEEK columns for high pressure liquid chromatography (HPLC). Briefly, plasma was depleted of the fourteen most abundant proteins using a multiple affinity removal system (MARS-14; Agilent). Depleted plasma was trypsin-digested and separated into flow-through and bound fractions by SNA or AAL HPLC. The fractions were treated with PNGaseF to remove N-linked glycans, and analyzed by LC-MS/MS on a QStar Elite. Data were analyzed using Mascot software. The experimental design included positive controls—fucosylated and sialylated human lactoferrin glycopeptides—and negative controls—high mannose glycopeptides from Saccharomyces cerevisiae—that were used to monitor the specificity of lectin capture. Key features of this workflow include the reproducibility derived from the HPLC format, the positive identification of the captured and PNGaseF-treated glycopeptides from their deamidated Asn-Xxx-Ser/Thr motifs, and quality assessment using glycoprotein standards. Protocol optimization also included determining the appropriate ratio of starting material to column capacity, identifying the most efficient capture and elution buffers, and monitoring the PNGaseF-treatment to ensure full deglycosylation. Future directions include using this workflow to perform mass spectrometry-based discovery experiments on plasma from breast cancer patients and control individuals.
Basic Protocols, Issue 32, Lectins, chromatography, glycopeptides, glycoproteins, biomarker discovery
Using Unfixed, Frozen Tissues to Study Natural Mucin Distribution
Institutions: University of California, San Diego , Los Alamos National Laboratory.
Mucins are complex and heavily glycosylated O
-linked glycoproteins, which contain more than 70% carbohydrate by weight1-3
. Secreted mucins, produced by goblet cells and the gastric mucosa, provide the scaffold for a micrometers-thick mucus layer that lines the epithelia of the gut and respiratory tract3,4
. In addition to mucins, mucus layers also contain antimicrobial peptides, cytokines, and immunoglobulins5-9
. The mucus layer is an important part of host innate immunity, and forms the first line of defense against invading microorganisms8,10-12
. As such, the mucus is subject to numerous interactions with microbes, both pathogens and symbionts, and secreted mucins form an important interface for these interactions. The study of such biological interactions usually involves histological methods for tissue collection and staining. The two most commonly used histological methods for tissue collection and preservation in the clinic and in research laboratories are: formalin fixation followed by paraffin embedding, and tissue freezing, followed by embedding in cryo-protectant media.
Paraffin-embedded tissue samples produce sections with optimal qualities for histological visualization including clarity and well-defined morphology. However, during the paraffin embedding process a number of epitopes become altered and in order to study these epitopes, tissue sections have to be further processed with one of many epitope retrieval methods13
. Secreted mucins and lipids are extracted from the tissue during the paraffin-embedding clearing step, which requires prolong incubation with organic solvents (xylene or Citrisolv). Therefore this approach is sub-optimal for studies focusing on the nature and distribution of mucins and mucus in vivo
In contrast, freezing tissues in Optimal Cutting Temperature (OCT) embedding medium avoids dehydration and clearing of the sample, and maintains the sample hydration. This allows for better preservation of the hydrated mucus layer, and thus permits the study of the numerous roles of mucins in epithelial biology. As this method requires minimal processing of the tissue, the tissue is preserved in a more natural state. Therefore frozen tissues sections do not require any additional processing prior to staining and can be readily analyzed using immunohistochemistry methods.
We demonstrate the preservation of micrometers-thick secreted mucus layer in frozen colon samples. This layer is drastically reduced when the same tissues are embedded in paraffin. We also demonstrate immunofluorescence staining of glycan epitopes presented on mucins using plant lectins. The advantage of this approach is that it does not require the use of special fixatives and allows utilizing frozen tissues that may already be preserved in the laboratory.
Medicine, Issue 67, Cellular Biology, Molecular Biology, Immunology, Biomedical Engineering, mucus, lectins, OCT, imaging, sialic acids, glycosylation
Improved In-gel Reductive β-Elimination for Comprehensive O-linked and Sulfo-glycomics by Mass Spectrometry
Institutions: University of Georgia, University of Georgia, Ishikawa Prefectural University.
Separation of proteins by SDS-PAGE followed by in-gel proteolytic digestion of resolved protein bands has produced high-resolution proteomic analysis of biological samples. Similar approaches, that would allow in-depth analysis of the glycans carried by glycoproteins resolved by SDS-PAGE, require special considerations in order to maximize recovery and sensitivity when using mass spectrometry (MS) as the detection method. A major hurdle to be overcome in achieving high-quality data is the removal of gel-derived contaminants that interfere with MS analysis. The sample workflow presented here is robust, efficient, and eliminates the need for in-line HPLC clean-up prior to MS. Gel pieces containing target proteins are washed in acetonitrile, water, and ethyl acetate to remove contaminants, including polymeric acrylamide fragments. O-linked glycans are released from target proteins by in-gel reductive β-elimination and recovered through robust, simple clean-up procedures. An advantage of this workflow is that it improves sensitivity for detecting and characterizing sulfated glycans. These procedures produce an efficient separation of sulfated permethylated glycans from non-sulfated (sialylated and neutral) permethylated glycans by a rapid phase-partition prior to MS analysis, and thereby enhance glycomic and sulfoglycomic analyses of glycoproteins resolved by SDS-PAGE.
Chemistry, Issue 93, glycoprotein, glycosylation, in-gel reductive β-elimination, O-linked glycan, sulfated glycan, mass spectrometry, protein ID, SDS-PAGE, glycomics, sulfoglycomics
Analysis of Nephron Composition and Function in the Adult Zebrafish Kidney
Institutions: University of Notre Dame.
The zebrafish model has emerged as a relevant system to study kidney development, regeneration and disease. Both the embryonic and adult zebrafish kidneys are composed of functional units known as nephrons, which are highly conserved with other vertebrates, including mammals. Research in zebrafish has recently demonstrated that two distinctive phenomena transpire after adult nephrons incur damage: first, there is robust regeneration within existing nephrons that replaces the destroyed tubule epithelial cells; second, entirely new nephrons are produced from renal progenitors in a process known as neonephrogenesis. In contrast, humans and other mammals seem to have only a limited ability for nephron epithelial regeneration. To date, the mechanisms responsible for these kidney regeneration phenomena remain poorly understood. Since adult zebrafish kidneys undergo both nephron epithelial regeneration and neonephrogenesis, they provide an outstanding experimental paradigm to study these events. Further, there is a wide range of genetic and pharmacological tools available in the zebrafish model that can be used to delineate the cellular and molecular mechanisms that regulate renal regeneration. One essential aspect of such research is the evaluation of nephron structure and function. This protocol describes a set of labeling techniques that can be used to gauge renal composition and test nephron functionality in the adult zebrafish kidney. Thus, these methods are widely applicable to the future phenotypic characterization of adult zebrafish kidney injury paradigms, which include but are not limited to, nephrotoxicant exposure regimes or genetic methods of targeted cell death such as the nitroreductase mediated cell ablation technique. Further, these methods could be used to study genetic perturbations in adult kidney formation and could also be applied to assess renal status during chronic disease modeling.
Cellular Biology, Issue 90,
zebrafish; kidney; nephron; nephrology; renal; regeneration; proximal tubule; distal tubule; segment; mesonephros; physiology; acute kidney injury (AKI)
Visualizing Neuroblast Cytokinesis During C. elegans Embryogenesis
Institutions: Concordia University.
This protocol describes the use of fluorescence microscopy to image dividing cells within developing Caenorhabditis elegans
embryos. In particular, this protocol focuses on how to image dividing neuroblasts, which are found underneath the epidermal cells and may be important for epidermal morphogenesis. Tissue formation is crucial for metazoan development and relies on external cues from neighboring tissues. C. elegans
is an excellent model organism to study tissue morphogenesis in vivo
due to its transparency and simple organization, making its tissues easy to study via microscopy. Ventral enclosure is the process where the ventral surface of the embryo is covered by a single layer of epithelial cells. This event is thought to be facilitated by the underlying neuroblasts, which provide chemical guidance cues to mediate migration of the overlying epithelial cells. However, the neuroblasts are highly proliferative and also may act as a mechanical substrate for the ventral epidermal cells. Studies using this experimental protocol could uncover the importance of intercellular communication during tissue formation, and could be used to reveal the roles of genes involved in cell division within developing tissues.
Neuroscience, Issue 85, C. elegans, morphogenesis, cytokinesis, neuroblasts, anillin, microscopy, cell division
A Method for Culturing Embryonic C. elegans Cells
Institutions: University of Miami .
is a powerful model system, in which genetic and molecular techniques are easily applicable. Until recently though, techniques that require direct access to cells and isolation of specific cell types, could not be applied in C. elegans
. This limitation was due to the fact that tissues are confined within a pressurized cuticle which is not easily digested by treatment with enzymes and/or detergents. Based on early pioneer work by Laird Bloom, Christensen and colleagues 1
developed a robust method for culturing C. elegans
embryonic cells in large scale. Eggs are isolated from gravid adults by treatment with bleach/NaOH and subsequently treated with chitinase to remove the eggshells. Embryonic cells are then dissociated by manual pipetting and plated onto substrate-covered glass in serum-enriched media. Within 24 hr of isolation cells begin to differentiate by changing morphology and by expressing cell specific markers. C. elegans
cells cultured using this method survive for up 2 weeks in vitro
and have been used for electrophysiological, immunochemical, and imaging analyses as well as they have been sorted and used for microarray profiling.
Developmental Biology, Issue 79, Eukaryota, Biological Phenomena, Cell Physiological Phenomena, C. elegans, cell culture, embryonic cells
A Manual Small Molecule Screen Approaching High-throughput Using Zebrafish Embryos
Institutions: University of Notre Dame.
Zebrafish have become a widely used model organism to investigate the mechanisms that underlie developmental biology and to study human disease pathology due to their considerable degree of genetic conservation with humans. Chemical genetics entails testing the effect that small molecules have on a biological process and is becoming a popular translational research method to identify therapeutic compounds. Zebrafish are specifically appealing to use for chemical genetics because of their ability to produce large clutches of transparent embryos, which are externally fertilized. Furthermore, zebrafish embryos can be easily drug treated by the simple addition of a compound to the embryo media. Using whole-mount in situ
hybridization (WISH), mRNA expression can be clearly visualized within zebrafish embryos. Together, using chemical genetics and WISH, the zebrafish becomes a potent whole organism context in which to determine the cellular and physiological effects of small molecules. Innovative advances have been made in technologies that utilize machine-based screening procedures, however for many labs such options are not accessible or remain cost-prohibitive. The protocol described here explains how to execute a manual high-throughput chemical genetic screen that requires basic resources and can be accomplished by a single individual or small team in an efficient period of time. Thus, this protocol provides a feasible strategy that can be implemented by research groups to perform chemical genetics in zebrafish, which can be useful for gaining fundamental insights into developmental processes, disease mechanisms, and to identify novel compounds and signaling pathways that have medically relevant applications.
Developmental Biology, Issue 93, zebrafish, chemical genetics, chemical screen, in vivo small molecule screen, drug discovery, whole mount in situ hybridization (WISH), high-throughput screening (HTS), high-content screening (HCS)
Flat Mount Preparation for Observation and Analysis of Zebrafish Embryo Specimens Stained by Whole Mount In situ Hybridization
Institutions: University of Notre Dame.
The zebrafish embryo is now commonly used for basic and biomedical research to investigate the genetic control of developmental processes and to model congenital abnormalities. During the first day of life, the zebrafish embryo progresses through many developmental stages including fertilization, cleavage, gastrulation, segmentation, and the organogenesis of structures such as the kidney, heart, and central nervous system. The anatomy of a young zebrafish embryo presents several challenges for the visualization and analysis of the tissues involved in many of these events because the embryo develops in association with a round yolk mass. Thus, for accurate analysis and imaging of experimental phenotypes in fixed embryonic specimens between the tailbud and 20 somite stage (10 and 19 hours post fertilization (hpf), respectively), such as those stained using whole mount in situ
hybridization (WISH), it is often desirable to remove the embryo from the yolk ball and to position it flat on a glass slide. However, performing a flat mount procedure can be tedious. Therefore, successful and efficient flat mount preparation is greatly facilitated through the visual demonstration of the dissection technique, and also helped by using reagents that assist in optimal tissue handling. Here, we provide our WISH protocol for one or two-color detection of gene expression in the zebrafish embryo, and demonstrate how the flat mounting procedure can be performed on this example of a stained fixed specimen. This flat mounting protocol is broadly applicable to the study of many embryonic structures that emerge during early zebrafish development, and can be implemented in conjunction with other staining methods performed on fixed embryo samples.
Developmental Biology, Issue 89, animals, vertebrates, fishes, zebrafish, growth and development, morphogenesis, embryonic and fetal development, organogenesis, natural science disciplines, embryo, whole mount in situ hybridization, flat mount, deyolking, imaging
Obtaining Eggs from Xenopus laevis Females
Institutions: Emory University.
The eggs of Xenopus laevis intact, lysed, and/or fractionated are useful for a wide variety of experiments. This protocol shows how to induce egg laying, collect and dejelly the eggs, and sort the eggs to remove any damaged eggs.
Basic Protocols, Issue 18, Current Protocols Wiley, Eggs, Xenopus laevis
Imaging Glycans in Zebrafish Embryos by Metabolic Labeling and Bioorthogonal Click Chemistry
Institutions: Albert Einstein College of Medicine, Yeshiva University, Albert Einstein College of Medicine, Yeshiva University, Albert Einstein College of Medicine, Yeshiva University.
Imaging glycans in vivo
has recently been enabled using a bioorthogonal chemical reporter strategy by treating cells or organisms with azide- or alkyne-tagged monosaccharides1, 2
. The modified monosaccharides, processed by the glycan biosynthetic machinery, are incorporated into cell surface glycoconjugates. The bioorthogonal azide or alkyne tags then allow covalent conjugation with fluorescent probes for visualization, or with affinity probes for enrichment and glycoproteomic analysis. This protocol describes the procedures typically used for noninvasive imaging of fucosylated glycans in zebrafish embryos, including: 1) microinjection of one-cell stage embryos with GDP-5-alkynylfucose (GDP-FucAl), 2) labeling fucosylated glycans in the enveloping layer of zebrafish embryos with azide-conjugated fluorophores via biocompatible Cu(I)-catalyzed azide-alkyne cycloaddition (CuAAC), and 3) imaging by confocal microscopy3
. The method described here can be readily extended to visualize other classes of glycans, e.g. glycans containing sialic acid4
, in developing zebrafish and in other living organisms.
Developmental Biology, Issue 52, click chemistry, chemical glycobiology, fucosylated glycans, embryogenesis, microinjection