This work was supported by NIH/NIDCR Grant 1R01DE019796.

The presented LeO-workflows allow for the stable isotope labeling of protein digests and synthetic peptides. These time course experiments (Figure 1) are applicable to comparative and quantitative proteomics studies as well as protease research. Each workflow consists of two experimental steps (Figure 2): A) The time resolved sampling of the respective 18O-stable isotope-encoded reaction (protease-catalyzed peptide cleavage; protease-catalyzed carboxyl oxygen exchange reaction; acid-catalyzed carboxyl oxygen exchange reaction) and B) analysis by mass spectrometry and graphical representation of 18O-incorporation kinetics.
A. TimeCourse Experiments
I. PALeO-TimeCourse: Protease-catalyzed labeling of proteolytic cleavages
<ol>
 <li>(Optional) Disulfide bonds of proteins (10 &mu;M) and peptides (250 &mu;M) are reduced with DTT (final concentration 2.5 mM) in 25 mM NH4HCO3 (both freshly prepared) by incubation for 30 min at 50 &deg;C.</li>
 <li>(Optional) Free cysteines are alkylated with iodoacetamide (final concentration 10 mM) in 25 mM NH4HCO3 by incubation for 30 min at room temperature in the dark.</li>
 <li>Depending on protease of interest, protein/peptide solutions need to be cleaned-up to remove residual buffer and alkylation agent. Use PepClean C-18 spin columns (Thermo) for peptide cleanup and Vivaspin Centrifugal Concentrators (Sartorius) to exchange buffers for protein samples.</li>
 <li>Redissolve/exchange peptides/proteins in 20 &mu;l protease reaction buffer (ECE-1: 50 mM MES-KOH, pH 5.5; trypsin: 25 mM NH4HCO3, pH 8.0) containing 1:1 (v/v) final H218O (95%, Sigma Isotec).</li>
 <li>Withdraw a zero time point sample prior to the addition of the protease: Mix 0.5 &mu;l of the reaction mixture and 0.5 &mu;l alpha cyano-4-hydroxycinnamic acid matrix (10 mg/ml in 50% acetonitrile, 0.1% TFA). Spot sample on an Opti-TOF 384 MALDI target plate (AB SCIEX) and leave the solvent droplets at room temperature until dry (about 5 min).</li>
 <li>Split the reaction solution into two aliquots. The first aliquot will be incubated with the protease of interest (ECE-1: 75 nM; trypsin: 0.04 nM) at room temperature or the recommended temperature for the particular enzyme. The second aliquot will be incubated without protease and will serve as control sample. The first aliquot represents a standard sample for protease-catalyzed 18O-labeling and can be used for peptide and protein identification, detection of proteolytic activities and monitoring of proteolytic cleavage reactions. The second aliquot is used to show that 18O-incorporation is induced by the protease; therefore, neither labeling nor cleavage is expected for this sample.</li>
 <li>Follow the reaction by removing reaction aliquots and spotting them as described under step 5 at time intervals as seem fit. The reaction conditions described above are used to monitor a proteolytic reaction for up to four days (approx. 24 spots). Start sampling every 5 min until 30 min, then spot every 30 min until 2 hr, then every hr until 8 hr and every 8 hr until 4 days. Cleavage products should appear within the first 12 hr and the substrate should be completely hydrolyzed after 24 hr. Extended reaction incubation times allow to define stable reaction products that are discrete from reaction intermediates, which are further processed. Depending on research question and given enzyme-substrate pairs, spotting times and incubation temperature have to be modulated to assure optimal reaction sampling.</li>
 <li>After the final reaction time point is spotted or in between extended spotting intervals, the MALDI target plate is submitted to MALDI-TOF/TOF MS/MS analysis as described below (Section B).</li>
</ol>
II. PALeO-TimeCourse: Postdigestion labeling of proteolytic termini
<ol>
 <li>(Optional) Disulfide bonds of proteins (10 &mu;M) and peptides (250 &mu;M) are reduced with DTT (final concentration 2.5 mM) in 25 mM NH4HCO3 (both freshly prepared) by incubation for 30 min at 50 &deg;C.</li>
 <li>(Optional) Free cysteines are alkylated with iodoacetamide (final concentration 10 mM) in 25 mM NH4HCO3 by incubation for 30 min at room temperature in the dark.</li>
 <li>Depending on the protease of interest, protein/peptide solutions need to be cleaned-up to remove residual buffer and alkylation agent. Use PepClean C-18 spin columns for peptide cleanup and Vivaspin Centrifugal Concentrators for protein buffer exchange.</li>
 <li>Redissolve/exchange the cleaned-up peptide products/proteins in 20 &mu;l protease reaction buffer (e.g. trypsin: 25 mM NH4HCO3, pH 8.0) and digest with protease of interest to completion (e.g. trypsin: 0.04 nM; 37 &deg;C; 12 hr).</li>
 <li>Cleanup cleavage products with PepClean C-18 spin columns. This step will eliminate residual protease activities.</li>
 <li>Redissolve/exchange the cleaned-up peptide products in 20 &mu;l protease reaction buffer (trypsin: 25 mM NH4HCO3, pH 8.0) containing 1:1 (v/v) final H218O.</li>
 <li>Withdraw a zero time point sample prior to the addition of enzyme: Mix 0.5 &mu;l of the reaction mixture and 0.5 &mu;l alpha cyano-4-hydroxycinnamic acid matrix (10 mg/ml in 50% acetonitrile, 0.1% TFA). Spot sample on a MALDI target plate and leave the solvent droplets at room temperature until dry (about 5 min).</li>
 <li>Split the reaction solution into two aliquots. The first aliquot will be incubated with protease of interest (e.g. trypsin: 0.04 nM) at room temperature or the recommended temperature for the particular enzyme. The second aliquot will be incubated without protease and will serve as a control. The first aliquot represents a standard sample for protease-catalyzed 18O-postdigestion labeling and can be used for peptide and protein quantification. The second aliquot is used to show that 18O-incorporation is catalyzed by the protease; therefore, no labeling is expected for this sample.</li>
 <li>Follow the reaction by removing reaction aliquots and spotting them as described under step 7.) at time intervals as seem fit. For example, we spotted initially every 5 min for up to 30 min and every 15 min after that to monitor the carboxyl oxygen exchange reaction catalyzed by trypsin.</li>
 <li>After the final reaction time point is spotted or in between extended spotting intervals the MALDI target plate is submitted to MALDI-TOF/TOF MS/MS analysis as described below (Section B).</li>
</ol>
III. ALeO-TimeCourse: Acid-catalyzed labeling of carboxyl groups
<ol>
 <li>Incubate individual peptides (50 nM) with 1:1 (v/v) 18O-enriched water in the presence or absence (control) of 0.1% (v/v) final trifluoroacetic acid (total volume 30 &mu;l).</li>
 <li>Sample the reaction products daily for 48 days by co-spotting a 0.5 &mu;l aliquot of the mixture with 0.5&mu;l of alpha cyano-4-hydroxycinnamic acid matrix (10 mg/ml in 50% acetonitrile, 0.1% TFA) directly onto a MALDI target plate.</li>
 <li>Between spotting intervals and after spotting of the final reaction time point submit MALDI target plate for MALDI-TOF/TOF MS/MS analysis as described below in Section B.</li>
</ol>
B. MALDI-TOF/TOF MS/MS Data Acquisition and Analysis
<ol>
 <li>Mass spectra are acquired on a 4800 MALDI TOF/TOF Analyzer (AB SCIEX).</li>
 <li>Prior to analysis, the instrument is calibrated with a mixture of peptide standards (Mass Standards Kit for Calibration of AB SCIEX TOF/TOF instruments) with a maximum mass measurement error tolerance of &plusmn; 50 ppm and a minimum number of six peaks to match.</li>
 <li>MS spectra (mass range 400 - 4,000 m/z) are acquired in triplicate using positive ion mode with an adjustable laser intensity (3,400 - 3,800; step size 50) with an acceptable base peak intensity range of 2,000 - 45,000. Single shots are acquired for sub-spectra, with 400 total shots/spectrum, stop conditions come into effect after 800 sub-spectra are acquired (pass or fail) or 400 sub-spectra pass acceptance criteria. In case of low-abundant samples or in the presence of complex biological backgrounds the lower end of the MS detection range should be raised to 800 m/z. In addition, it may be necessary to remove salt and other interfering compounds with a sample cleanup step as described earlier or by LC separation.</li>
 <li>MS data files (.t2d files) are exported from the 4000 Series Explorer data acquisition software and imported into our in-house laboratory information system, which utilizes MASCOT Distiller software (Matrix Science) for spectral processing and peak detection. Isotopic envelopes are deconvoluted and 18O-incorporation ratios automatically determined using an algorithm similar to the one described by Mason et al.12 and adapted by our group9. Alternatively, software tools such as ZoomQuant13 and Viper14, as well as commercial software packages such as BioWorks Xpress (Thermo Fisher Scientific) and Mascot Distiller Quantitation Toolbox (Matrix Science) can deconvolute 18O-type data15,16. 18O-incorporation ratios are expressed as the relative contributions of individual peptide isotope species (i.e., peptides containing 16O, 18O1 or 18O2) to the entire isotopic envelope.</li>
 <li>For each TimeCourse experiment, the molecular masses ([M+H]+) for all detected peptide species are extracted from the associated MS data files and the values binned at a 100 ppm mass width.</li>
 <li>At least three [M+H]+ are set to be required to populate a bin. In case of known substrates, the filtered bin list is compared to a list of proteolytic cleavage products predicted from the substrate peptide sequence using the ExPASy FindPept tool17 (<a href="http://au.expasy.org/tools/findpept.html" target="_blank">http://au.expasy.org/tools/findpept.html</a>) and a 200 ppm mass error acceptance tolerance.</li>
 <li>MS/MS spectra are acquired for all mass values of cleavage products predicted by the FindPept tool and for bin values that have 18O-incorporations associated with them. MS/MS data are acquired on the 4800 MALDI TOF/TOF Analyzer in 1kV reflector positive ion mode, with a fixed laser intensity of 4200 and CID-gas in off mode. 50 shots are acquired in a randomized pattern per sub-spectra up to a total of 40 sub-spectra per spot (yielding a total of 2,000 shots/spot).</li>
 <li>MS/MS data files (.t2d files) are exported from the 4000 Series Explorer data acquisition software and imported into our in-house laboratory information system, peaks are detected and MS/MS peak lists are associated with the corresponding binned MS data.</li>
 <li>For peptide identification, MS/MS peak lists are searched against the SwissProt database using the MASCOT search engine with the following search parameters: no enzyme specificity, 150 ppm precursor ion and 0.2 Da fragment ion mass tolerances.</li>
 <li>Peptide identifications can additionally be validated using the Data Explorer software (AB SCIEX) by confirming the characteristic 18O-incorporation patterns across y-series fragment ions as described by Shevchenko et al.18.</li>
</ol>
C. Preparation of Spectral Time and 18O-incorporation Plots
Spectral time plots: MS data files (.t2d files) for each reaction time point are exported from the Data Explorer software as ASCII-files using a macro and imported into a data analysis and graphic software program (e.g. Origin by OriginLab) and displayed as waterfall plots (Figure 3).
18O-incorporation plots: For each binned peptide cleavage product, the relative contributions of individual peptide isotope species (16O, 18O1 or 18O2) are extracted across all reaction time points and plotted against time (Figure 4).

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No conflicts of interest declared.

By combining stable isotope labeling and high-resolution mass spectrometry in a time-resolved manner, the PALeO-TimeCourse method allows for a dynamic analysis of the generation of peptide products. The assay can be used to generate stable isotopically labeled peptides for quantitative and qualitative proteomics studies and to evaluate the kinetics by which proteotypic peptides are generated. Furthermore, PALeO-TimeCourse is designed to evaluate proteolytic pathways under specific, physiologically relevant conditions <em...

<table border="1">
 <tbody>
 <tr>
 <td>Name of material</td>
 <td>Company</td>
 <td>Catalogue number</td>
 </tr>
 <tr>
 <td>PepClean C-18 Spin Columns</td>
 <td>Thermo</td>
 <td>89870</td>
 </tr>
 <tr>
 <td>Opti-TOF 384 MALDI target plate </td>
 <td>AB SCIEX </td>
 <td>1016629</td>
 </tr>
 <tr>
 <td>4800 MALDI TOF/TOF</td>
 <td>AB SCIEX </td>
 <td></td>
 </tr>
 </tbody>
</table>
Table 1. Materials
<table border="1">
 <tbody>
 <tr>
 <td>Name of reagent</td>
 <td>Company</td>
 <td>Catalogue number</td>
 </tr>
 <tr>
 <td>Alpha cyano-4-hydroxycinnamic acid</td>
 <td>Sigma Aldrich</td>
 <td>70990-1G-F</td>
 </tr>
 <tr>
 <td>Bovine serum albumin (BSA)</td>
 <td>Sigma Aldrich</td>
 <td>A3294-10G</td>
 </tr>
 <tr>
 <td>Dithiothreitol (DTT)</td>
 <td>Acros</td>
 <td>16568-0050</td>
 </tr>
 <tr>
 <td>Iodoacetamide (IAM)</td>
 <td>Sigma Aldrich</td>
 <td>1149-5G</td>
 </tr>
 <tr>
 <td>Endothelin converting enzyme-1 (ECE-1)</td>
 <td>R&amp;D Systems</td>
 <td>1784-ZN</td>
 </tr>
 <tr>
 <td>Trypsin Gold</td>
 <td>Promega</td>
 <td>V5280</td>
 </tr>
 <tr>
 <td>Water-18O, 97 atom % 18O</td>
 <td>Sigma Aldrich</td>
 <td>329878-1G</td>
 </tr>
 <tr>
 <td>Trifluoroacetic acid (TFA)</td>
 <td>Thermo</td>
 <td>28904</td>
 </tr>
 <tr>
 <td>Mass Standards Kit for Calibration of AB SCIEX TOF/TOF instruments</td>
 <td>AB SCIEX</td>
 <td>4333604</td>
 </tr>
 </tbody>
</table>
Table 2. Reagents

protease- and acid-catalyzed labeling workflows employing 18o-enriched water

Stable isotopes are essential tools in biological mass spectrometry. Historically, 18O-stable isotopes have been extensively used to study the catalytic mechanisms of proteolytic enzymes1-3. With the advent of mass spectrometry-based proteomics, the enzymatically-catalyzed incorporation of 18O-atoms from stable isotopically enriched water has become a popular method to quantitatively compare protein expression levels (reviewed by Fenselau and Yao4, Miyagi and Rao5 and Ye et al.6). 18O-labeling constitutes a simple and low-cost alternative to chemical (e.g. iTRAQ, ICAT) and metabolic (e.g. SILAC) labeling techniques7. Depending on the protease utilized, 18O-labeling can result in the incorporation of up to two 18O-atoms in the C-terminal carboxyl group of the cleavage product3. The labeling reaction can be subdivided into two independent processes, the peptide bond cleavage and the carboxyl oxygen exchange reaction8. In our PALeO (protease-assisted labeling employing 18O-enriched water) adaptation of enzymatic 18O-labeling, we utilized 50% 18O-enriched water to yield distinctive isotope signatures. In combination with high-resolution matrix-assisted laser desorption ionization time-of-flight tandem mass spectrometry (MALDI-TOF/TOF MS/MS), the characteristic isotope envelopes can be used to identify cleavage products with a high level of specificity. We previously have used the PALeO-methodology to detect and characterize endogenous proteases9 and monitor proteolytic reactions10-11. Since PALeO encodes the very essence of the proteolytic cleavage reaction, the experimental setup is simple and biochemical enrichment steps of cleavage products can be circumvented. The PALeO-method can easily be extended to (i) time course experiments that monitor the dynamics of proteolytic cleavage reactions and (ii) the analysis of proteolysis in complex biological samples that represent physiological conditions. PALeO-TimeCourse experiments help identifying rate-limiting processing steps and reaction intermediates in complex proteolytic pathway reactions. Furthermore, the PALeO-reaction allows us to identify proteolytic enzymes such as the serine protease trypsin that is capable to rebind its cleavage products and catalyze the incorporation of a second 18O-atom. Such "double-labeling" enzymes can be used for postdigestion 18O-labeling, in which peptides are exclusively labeled by the carboxyl oxygen exchange reaction. Our third strategy extends labeling employing 18O-enriched water beyond enzymes and uses acidic pH conditions to introduce 18O-stable isotope signatures into peptides.

We used the PALeO-TimeCourse workflow to dynamically monitor the incorporation of 18O-stable isotopes into peptide cleavage products generated by proteolytic enzymes. The presented approach is a versatile tool to comparatively study proteolytic processing pathways for different substrate and protease combinations. By sampling proteolytic reactions repeatedly over the course of the reaction, the PALeO-TimeCourse experiment provides time-resolved snapshots of substrate and product abundances and processing detai...

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Protease- and Acid-catalyzed Labeling Workflows Employing 18O-enriched Water

Stable isotope labeling workflows employing 18O-enriched water (LeO-workflows) are versatile tools for quantitative and qualitative proteomics studies. In protease-assisted (PALeO) workflows, 18O-atoms are introduced by proteolytic cleavage and carboxyl oxygen exchange reactions mediated by proteases. In the acid-catalyzed (ALeO) workflow, 18O-atoms are introduced by carboxyl oxygen exchange at low pH.

Protease- and Acid-catalyzed Labeling Workflows Employing 18O-enriched Water

Stable isotopes are essential tools in biological mass spectrometry. Historically, 18O-stable isotopes have been extensively used to study the catalytic ...

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11.9K Views.  Boston Biomedical Research Institute. Stable isotope labeling workflows employing 18O-enriched water (LeO-workflows) are versatile tools for quantitative and qualitative proteomics studies. In protease-assisted (PALeO) workflows, 18O-atoms are introduced by proteolytic cleavage and carboxyl oxygen exchange reactions mediated by proteases. In the acid-catalyzed (ALeO) workflow, 18O-atoms are introduced by carboxyl oxygen exchange at low pH.

Video: Protease- and Acid-catalyzed Labeling Workflows Employing 18O-enriched Water

Measuring Exocytosis in Neurons Using FM Labeling

The ability to measure the kinetics of vesicle release can help provide insight into some of the basics of neurotransmission.  Here we used real-time imaging of vesicles labeled with FM dye to monitor the rate of presynaptic vesicle release.  FM4-64 is a red fluorescent amphiphilic styryl dye that embeds into the membranes of synaptic vesicles as endocytosis is stimulated.  Lipophilic interactions cause the dye to greatly increase in fluorescence, thus emitting a bright signal when associated with vesicles and a nominal one when in the extracellular fluid. After a wash step is used to help remove external dye within the plasma membrane, the remaining FM is concentrated within the vesicles and is then expelled when exocytosis is induced by another round of electrical stimulation. The rate of vesicles release is measured from the resulting decrease in fluorescence.  Since FM dye can be applied external and transiently, it is a useful tool for determining rates of exocytosis in neuronal cultures, especially when comparing the rates between transfected synapses and neighboring control boutons.



The ability to measure the kinetics of vesicle release can help provide insight into some of the basics of neurotransmission. Here we used real-time imaging of vesicles labeled with the red fluorescent dye FM 4-64 to measure the rate of presynaptic vesicle release in hippocampal neuronal cultures.

Pomiar egzocytozy w Neurony Korzystanie FM etykietowania

The ability to measure the kinetics of vesicle release can help provide insight into some of the basics of neurotransmission.  Here we used real-time ...

Morris Water Maze Experiment

The Morris water maze is widely used to study spatial memory and learning. Animals are placed in a pool of water that is colored opaque with powdered non-fat milk or non-toxic tempera paint, where they must swim to a hidden escape platform. Because they are in opaque water, the animals cannot see the platform, and cannot rely on scent to find the escape route. Instead, they must rely on external/extra-maze cues. As the animals become more familiar with the task, they are able to find the platform more quickly. Developed by Richard G. Morris in 1984, this paradigm has become one of the "gold standards" of behavioral neuroscience.

The Morris water maze is a well-accepted tool used to document the involvement of the hippocampus in a behavioral task.

The Morris water maze is widely used to study spatial memory and learning. Animals are placed in a pool of water that is colored opaque with powdered ...

Sigma's Non-specific Protease Activity Assay - Casein as a Substrate

Proteases break peptide bonds. In the lab, it is often necessary to measure and/or compare the activity of proteases. Sigma's non-specific protease activity assay may be used as a standardized procedure to determine the activity of proteases, which is what we do during our quality control procedures. In this assay, casein acts as a substrate. When the protease we are testing digests casein, the amino acid tyrosine is liberated along with other amino acids and peptide fragments. Folin and Ciocalteus Phenol, or Folin's reagent primarily reacts with free tyrosine to produce a blue colored chromophore, which is quantifiable and measured as an absorbance value on the spectrophotometer. The more tyrosine that is released from casein, the more the chromophores are generated and the stronger the activity of the protease. Absorbance values generated by the activity of the protease are compared to a standard curve, which is generated by reacting known quantities of tyrosine with the F-C reagent to correlate changes in absorbance with the amount of tyrosine in micromoles. From the standard curve the activity of protease samples can be determined in terms of Units, which is the amount in micromoles of tyrosine equivalents released from casein per minute.
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Proteases break peptide bonds. In the lab, it is often necessary to measure and/or compare the activity of proteases. Sigma's non-specific protease activity assay may be used as a standardized procedure to determine the activity of proteases.

Proteases break peptide bonds. In the lab, it is often necessary to measure and/or compare the activity of proteases.  Sigma's non-specific protease ...

Fluorescent Labeling of Drosophila Heart Structures

The Drosophila melanogaster dorsal vessel, or heart, is a tubular structure comprised of a single layer of contractile cardiomyocytes, pericardial cells that align along each side of the heart wall, supportive alary muscles and, in adults, a layer of ventral longitudinal muscle cells. The contractile fibers house conserved constituents of the muscle cytoarchitecture including densely packed bundles of myofibrils and cytoskeletal/submembranous protein complexes, which interact with homologous components of the extracellular matrix. Here we describe a protocol for the fixation and the fluorescent labeling of particular myocardial elements from the hearts of dissected larvae and semi-intact adult Drosophila. Specifically, we demonstrate the labeling of sarcomeric F-actin and of &alpha;-actinin in larval hearts. Additionally, we perform labeling of F-actin and &alpha;-actinin in myosin-GFP expressing adult flies and of &alpha;-actinin and pericardin, a type IV extracellular matrix collagen, in wild type adult hearts. Particular attention is given to a mounting strategy for semi-intact adult hearts that minimizes handling and optimizes the opportunity for maintaining the integrity of the cardiac tubes and the associated tissues. These preparations are suitable for imaging via fluorescent and confocal microscopy. Overall, this procedure allows for careful and detailed analysis of the structural characteristics of the heart from a powerful genetically tractable model system.

Fluorescent Labeling of Drosophila Heart Structures

Here we describe a basic protocol for fluorescent labeling of different elements of heart tubes from larva and adult Drosophila melanogaster. These specimens are well-suited for imaging via fluorescent or confocal microscopy. This technique permits detailed structural analysis of the features of the hearts from a powerful model organism.

The Drosophila melanogaster dorsal vessel, or heart, is a tubular structure comprised of a single layer of contractile cardiomyocytes, pericardial cells ...

Use of the Protease Fluorescent Detection Kit to Determine Protease Activity

The Protease Fluorescent Detection Kit provides ready-to-use reagents for detecting the presence of protease activity. This simple assay to detect protease activity uses casein labeled with fluorescein isothiocyanate (FITC) as the substrate.
Protease activity results in the cleavage of the FITC-labeled casein substrate into smaller fragments, which do not precipitate under acidic conditions. After incubation of the protease sample and substrate, the reaction is acidified with the addition of trichloroacetic acid (TCA). The mixture is then centrifuged with the undigested substrate forming a pellet and the smaller, acid soluble fragments remaining in solution. The supernatant is neutralized and the fluorescence of the FITC-labeled fragments is measured.
The described kit procedure detects the trypsin protease control at a concentration of approximately 0.5 &mu;g/ml (5 ng of trypsin added to the assay). This sensitivity can be increased with a longer incubation time, up to 24 hours. The assay is performed in microcentrifuge tubes and procedures are provided for fluorescence detection using either cuvettes or multiwell plates.

The Protease Fluorescent Detection Kit is designed for the measurement of protease activity using fluorometry. It is also suitable for detection of trace amounts of protease contamination. The method is based on the proteolytic hydroysis of a proprietary formulation of a FITC-labeled casein substrate.

The Protease Fluorescent Detection Kit provides ready-to-use reagents for detecting the presence of protease activity. This simple assay to detect ...

Preparation of Drosophila Polytene Chromosome Squashes for Antibody Labeling

Drosophila has long been a favorite model system for studying the relationship between chromatin structure and gene regulation due to the cytological advantages provided by the giant salivary gland polytene chromosomes of third instar larvae.  In this tissue the chromosomes undergo many rounds of replication in the absence of cell division giving rise to approximately 1000 copies.  The DNA remains aligned after each replicative cycle resulting in greatly enlarged chromosomes that provide a unique opportunity to correlate chromatin morphology with the localization of specific proteins.  Consequently, there has been a high level of interest in defining the epigenetic modifications present at different genes and at different stages of the transcription process. An important tool for such studies is the labeling of polytene chromosomes with antibodies to the enzyme, transcription factor, or histone modification of interest. This video protocol illustrates the squash technique used in the Johansen laboratory to prepare Drosophila polytene chromosomes for antibody labeling.

This video protocol illustrates the squash technique used in the Johansen laboratory to prepare Drosophila polytene chromosomes for antibody labeling.

Drosophila has long been a favorite model system for studying the relationship between chromatin structure and gene regulation due to the cytological ...

ReAsH/FlAsH Labeling and Image Analysis of Tetracysteine Sensor Proteins in Cells

Fluorescent proteins and dyes are essential tools for the study of protein trafficking, localization and function in cells. While fluorescent 
proteins such as green fluorescence protein (GFP) have been extensively used as fusion partners to proteins to track the properties of a protein of interest1, recent 
developments with smaller tags enable new functionalities of proteins to be examined in cells such as conformational change and protein-association 2, 3. One small 
tag system involves a tetracysteine motif (CCXXCC) genetically inserted into a target protein, which binds to biarsenical dyes, ReAsH (red fluorescent) and FlAsH 
(green fluorescent), with high specificity even in live cells 2. The TC/biarsenical dye system offers far less steric constraints to the host protein than 
fluorescent proteins which has enabled several new approaches to measure conformational change and protein-protein interactions 4-7. We recently developed 
a novel application of TC tags as sensors of oligomerization in cells expressing mutant huntingtin, which when mutated aggregates in neurons in Huntington 
disease 7. Huntingtin was tagged with two fluorescent dyes, one a fluorescent protein to track protein location, and the second a TC tag which only 
binds biarsenical dyes in monomers. Hence, changes in colocalization between protein and biarsenical dye reactivity enabled submicroscopic oligomer 
content to be spatially mapped within cells. Here, we describe how to label TC-tagged proteins fused to a fluorescent protein (Cherry, GFP or CFP) 
with FlAsH or ReAsH in live mammalian cells and how to quantify the two color fluorescence (Cherry/FlAsH, CFP/FlAsH or GFP/ReAsH combinations).

The biarsenical dyes FlAsH and ReAsH bind specifically to tetracysteine motifs in proteins and can selectively label proteins in live cells. Recently this labeling strategy has been used to develop sensors for different protein conformations or oligomeric states. We describe the labeling approach and methods to quantitatively analyze binding.

Fluorescent proteins and dyes are essential tools for the study of protein trafficking, localization and function in cells.  While fluorescent 
proteins ...

GC-based Detection of Aldononitrile Acetate Derivatized Glucosamine and Muramic Acid for Microbial Residue Determination in Soil

Quantitative approaches to characterizing microorganisms are crucial for a broader understanding of the microbial status and function within ecosystems. Current strategies for microbial analysis include both traditional laboratory culture-dependent techniques and those based on direct extraction and determination of certain biomarkers1, 2. Few among the diversity of microbial species inhabiting soil can be cultured, so culture-dependent methods introduce significant biases, a limitation absent in biomarker analysis.
The glucosamine, mannosamine, galactosamine and muramic acid have been well served as measures of both the living and dead microbial mass, of these the glucosamine (most abundant) and muramic acid (uniquely from bacterial cell) are most important constituents in the soil systems3, 4. However, the lack of knowledge on the analysis restricts the wide popularization among scientific peers. Among all existing analytical methods, derivatization to aldononitrile acetates followed by GC-based analysis has emerged as a good option with respect to optimally balancing precision, sensitivity, simplicity, good chromatographic separation, and stability upon sample storage5.
Here, we present a detailed protocol for a reliable and relatively simple analysis of glucosamine and muramic acid from soil after their conversion to aldononitrile acetates. The protocol mainly comprises four steps: acid digestion, sample purification, derivatization and GC determination. The step-by-step procedure is modified according to former publications6, 7. In addition, we present a strategy to structurally validate the molecular ion of the derivative and its ion fragments formed upon electron ionization. We applied GC-EI-MS-SIM, LC-ESI-TOF-MS and isotopically labeled reagents to determine the molecular weight of aldononitrile acetate derivatized glucosamine and muramic acid; we used the mass shift of isotope-labeled derivatives in the ion spectrum to investigate ion fragments of each derivatives8. In addition to the theoretical elucidation, the validation of molecular ion of the derivative and its ion fragments will be useful to researchers using &delta;13C or ion fragments of these biomarkers in biogeochemical studies9, 10.

We describe a method protocol for the GC-based analysis of the aldonitrile acetate derivatives of glucosamine and muramic acid extracted from soil. For elucidation of the chemical mechanism, we also present a strategy to confirm the structure of the derivative and the ion fragments formed upon electron ionization.

Quantitative approaches to characterizing microorganisms are crucial for a broader understanding of the microbial status and function within ecosystems. ...

Measuring Cell Cycle Progression Kinetics with Metabolic Labeling and Flow Cytometry

Precise control of the initiation and subsequent progression through the various phases of the cell cycle are of paramount importance in proliferating cells. Cell cycle division is an integral part of growth and reproduction and deregulation of key cell cycle components have been implicated in the precipitating events of carcinogenesis 1,2. Molecular agents in anti-cancer therapies frequently target biological pathways responsible for the regulation and coordination of cell cycle division 3. Although cell cycle kinetics tend to vary according to cell type, the distribution of cells amongst the four stages of the cell cycle is rather consistent within a particular cell line due to the consistent pattern of mitogen and growth factor expression. Genotoxic events and other cellular stressors can result in a temporary block of cell cycle progression, resulting in arrest or a temporary pause in a particular cell cycle phase to allow for instigation of the appropriate response mechanism. 
The ability to experimentally observe the behavior of a cell population with reference to their cell cycle progression stage is an important advance in cell biology. Common procedures such as mitotic shake off, differential centrifugation or flow cytometry-based sorting are used to isolate cells at specific stages of the cell cycle 4-6. These fractionated, cell cycle phase-enriched populations are then subjected to experimental treatments. Yield, purity and viability of the separated fractions can often be compromised using these physical separation methods. As well, the time lapse between separation of the cell populations and the start of experimental treatment, whereby the fractionated cells can progress from the selected cell cycle stage, can pose significant challenges in the successful implementation and interpretation of these experiments.
Other approaches to study cell cycle stages include the use of chemicals to synchronize cells. Treatment of cells with chemical inhibitors of key metabolic processes for each cell cycle stage are useful in blocking the progression of the cell cycle to the next stage. For example, the ribonucleotide reductase inhibitor hydroxyurea halts cells at the G1/S juncture by limiting the supply of deoxynucleotides, the building blocks of DNA. Other notable chemicals include treatment with aphidicolin, a polymerase alpha inhibitor for G1 arrest, treatment with colchicine and nocodazole, both of which interfere with mitotic spindle formation to halt cells in M phase and finally, treatment with the DNA chain terminator 5-fluorodeoxyridine to initiate S phase arrest 7-9. Treatment with these chemicals is an effective means of synchronizing an entire population of cells at a particular phase. With removal of the chemical, cells rejoin the cell cycle in unison. Treatment of the test agent following release from the cell cycle blocking chemical ensures that the drug response elicited is from a uniform, cell cycle stage-specific population. However, since many of the chemical synchronizers are known genotoxic compounds, teasing apart the participation of various response pathways (to the synchronizers vs. the test agents) is challenging. 
Here we describe a metabolic labeling method for following a subpopulation of actively cycling cells through their progression from the DNA replication phase, through to the division and separation of their daughter cells. Coupled with flow cytometry quantification, this protocol enables for measurement of kinetic progression of the cell cycle in the absence of either mechanically- or chemically- induced cellular stresses commonly associated with other cell cycle synchronization methodologies 10. In the following sections we will discuss the methodology, as well as some of its applications in biomedical research.

Tracking subtle changes in the progression and kinetics of cell cycle stages can be accomplished by use of a combination of metabolic labeling of nucleic acids with BrdU and total genomic DNA staining via Propidium Iodide. This method avoids the need of chemical synchronization of cycling cells, thereby preventing the introduction of non-specific DNA damage, which in turn affects cell cycle progression.

Precise control of the initiation and subsequent progression through the various phases of the cell cycle are of paramount importance in proliferating ...

Mouse Islet of Langerhans Isolation using a Combination of Purified Collagenase and Neutral Protease

The interrogation of beta cell gene expression and function in vitro has squarely shifted over the years from the study of rodent tumorigenic cell lines to the study of isolated rodent islets. Primary islets offer the distinct advantage that they more faithfully reflect the biology of intracellular signaling pathways and secretory responses. Whereas the method of islet isolation using tissue dissociating enzyme (TDE) preparations has been well established in many laboratories1-4, variations in the consistency of islet yield and quality from any given rodent strain limit the extent and feasibility of primary islet studies. These variations often occur as a result of the crude partially purified TDEs used in the islet isolation procedure; TDEs frequently exhibit lot-to-lot variations in activity and often require adjustments to the dose of enzyme used. A small number of reports have used purified TDEs for rodent cell isolations5, 6, but the practice is not widespread despite the routine use and advantages of purified TDEs for human islet isolations. In collaboration with VitaCyte, LLC (Indianapolis, IN), we developed a modified mouse islet isolation protocol based on that described by Gotoh7, 8, in which the TDEs are perfused directly into the pancreatic duct of mice, followed by crude tissue fractionation through a Histopaque gradient9, and isolation of purified islets. A significant difference in our protocol is the use of purified collagenase (CIzyme MA) and neutral protease (CIzyme BP) combination. The collagenase was characterized by the use of a6 fluorescence collagen degrading activity (CDA) assay that utilized fluorescently labeled soluble calf skin fibrils as substrate6. This substrate is more predictive of the kinetics of collagen degradation in the tissue matrix because it relies on native collagen as the substrate. The protease was characterized with a sensitive fluorescent kinetic assay10. Utilizing these improved assays along with more traditional biochemical analysis enable the TDE to be manufactured more consistently, leading to improved performance consistency between lots. The protocol described in here was optimized for maximal islet yield and optimal islet morphology using C57BL/6 mice. During the development of this protocol, several combinations of collagenase and neutral proteases were evaluated at different concentrations, and the final ratio of collagenase:neutral protease of 35:10 represents enzyme performance comparable to Sigma Type XI. Because significant variability in average islet yields from different strains of rats and mice have been reported, additional modifications of the TDE composition should be made to improve the yield and quality of islets recovered from different species and strains.

A detailed description of mouse islet isolation is described using the technique of in situ pancreatic ductal cannulation and perfusion of a combination of purified collagenase and neutral protease.

The interrogation of beta cell gene expression and function in vitro has squarely shifted over the years from the study of rodent tumorigenic cell lines ...