This work is funded by the DFG.

Bee Preparation and Back-fill
<ol>
<li>Catch honey bee foragers at hive and chill on ice for immobilization.</li>
<li>Mount in Plexiglas recording chambers3. Fix eyes and thorax to the walls of the recording chamber with low melting point hard wax.</li>
<li>Pull glass capillaries as commonly used for electrophysiology and break at the tip to obtain a tip diameter of ca. 10&mu;m. Cover tip of the capillary with dye paste. Dye paste consists of a 10:1 mixture of Fura-2 dextran and lysine fixable tetramethylrhodamine dextran.</li>
<li>For the staining procedure immobilize antennae with minutiae or n-eicosane. Remove a piece of cuticle above the brain and push glands and trachea to the side to allow access to the mushroom body.</li>
<li>Inject capillaries into either the soma or axonal region of mushroom body neurons. This can be done free hand or using a micro manipulator. Then restore cuticle piece on head capsule and loosen antennae.</li>
<li>Feed bees with 30% sucrose solution before storing them in a humidified case for at least four hours or over night at 20&deg;C.</li>
</ol>
In vivo Imaging
<ol>
<li>Prevent movements of the bee e.g. with a small piece of sponge that is pressed against the abdomen and fixed with a clip or tape to the recording chamber.</li>
<li>To prevent movement of the brain resulting from pumping of the oesophagus, cut a small incision into the cuticle above the labrum and pull out carefully the oesophagus and its surrounding solid structures to put it under tension without damaging the oesophagus4. Cover with two component silicone.</li>
<li>Make holes with a needle and insert wire electrodes for the recording of M17.</li>
<li>Remove cuticle piece, trachea and glands above the brain. Suck heamolymph inside the head capsule with a piece of paper.</li>
<li>Fill the head capsule with two component silicone. It is important that the brain is completely covered.</li>
<li>Place bee on microscope stage and place a drop of water on the surface of the silicone to immerse the dip objective of the microscope into the droplet. Focus on stained neurons.</li>
</ol>
Odor stimulation and recording of signals
Olfactometer: We use a computer controlled, custom built stimulation device or &ldquo;olfactometer&rdquo; as described previously by Galizia and Vetter3. Odors are diluted into a constant air stream directed towards the antennae. Any stimulus consists of a 3s odor pulse of 0.2ml odor saturated air. The stimulation protocol can be set in the recording software TILLVision. Microscope and Imaging set-up: We use a Zeiss fluorescence microscope. Images are recorded at 25&deg;C with a sampling-rate of 5 Hz using a TILL-Photonics imaging device mounted on the microscope. Calcium signals are recorded through an X60, 0.9 W Olympus dip objective with an Imago CCD camera (640X480 pixels, 4X binned on chip to 160X120 pixels). Fura-2 is excited with monochromatic light of 340 and 380nm wavelength for ratiometric measurements. Fluorescence is detected through a 410nm dichroic mirror and a 440nm long pass filter. The parameters for the recording are set in the recording software. Each measurement lasts 10s and exposure time for the two wavelengths can be adjusted to the staining intensity in each preparation. M17 recording: The protractor muscle of the labium (M17) is recorded extracellularly to monitor the behavioral responses related to learning, i.e. the PER 5, 6. Inject a copper wire into the muscle close to the mouth parts. Inject ground electrode into the eye. The muscle potentials are amplified with a custom build pre-amplifier, digitalized and stored on a computer. Stimulus onset is triggered by the olfactometer.
Data Analysis
<ol>
<li>During the experiment: The recordings of the calcium signal are fed into a computer and stored. The recording software TILLVision allows an initial inspection of the signals, like calculation of the ratio between the wavelengths (340nm/380nm) and calculation of deltaF (subtraction of baseline).</li>
<li>Image processing is performed with custom written programs in IDL. Calculate ratio of Ca2+ signals from 340nm and 380nm measurements for each pixel. Determine background fluorescence (F) by averaging over frames before the stimulus and subtract from the ratiometric signal (deltaF).</li>
<li>To define Region of Interest (ROI): Calculate mean of frames during the stimulus and apply a low pass filter (3X3px). Convert grey scale to false color scale. Determine activity in neuronal structures as activity spots in false color images and define as ROI.</li>
<li>Calculate temporal dynamics of the active regions by averaging the pixels in the ROI over all frames without any filtering or correction.</li>
</ol>
Morphological Analysis and Reconstruction
Together with the Fura-2 dextran we fill the neurons with the fixable dye rhodamine dextran (&ldquo;mini ruby&rdquo;). Both dyes have the same molecular weight; they should be co-located in the neurons. Wide-field imaging allows only very limited spatial resolution, therefore it is recommended to investigate the stained structures after the experiment using confocal scanning microscopy.
<ol>
<li>After physiological measurements, dissect brains and fixate.</li>
<li>Rinse in PBS and stepwise dehydrate in ethanol and clear in methyl salycilate.</li>
<li>For confocal microscopy embed brain in methyl salycilate.</li>
<li>We use a Leica TCS SP2 microscope. Excitation wavelength is 543nm.</li>
<li>We scan the brain with an air or oil immersion objective and compare the stained structures with the imaging data. </li>
</ol>
Representative Results:
If the staining and preparation was successful, the signal in false color scale, for example in mushroom body alpha lobe extrinsic neurons, could look like in figure 1. The behavioral response (PER) can be calculated as spike frequency during the stimulus minus spike frequency before the stimulus (figure2). The stained structures may be investigated in greater detail using a confocal microscope (figure3).
<img src="/files/ftp_upload/1353/figure1_new.png" alt="Figure 1" /> Figure 1: Representative Calcium Imaging signals (stimulation with heptanal). A: Imaged region of the bee brain. B: Raw fluorescent image as seen through a X20 dip objective at 380nm excitation wavelength, the alpha-lobe of the mushroom body is outlined in purple. Dye was injected into the ventro-lateral rim of the alpha-lobes to stain extrinsic neurons. C: False color image derived from the average signal, deltaF (340/380), during the three second stimulus; alpha-lobe outlined in purple, selected ROI outlined in black. D: Temporal dynamics of calcium signal in response to odor stimulation, average signal from region of interest as indicated in C. E: Temporal dynamics of calcium signal in response to sucrose stimulation of the ipsi- and contra-lateral antenna respectively to the ROI indicated in C.
<img src="/files/ftp_upload/1353/figure2_new.png" alt="" /> Figure 2: Representative M17 response. After olfactory conditioning the bee extends the proboscis when the odor is presented without reward. Red line indicates odor stimulus. Inset: PER of the honeybee.
<img src="/files/ftp_upload/1353/figure3_new.png" alt="" /> Figure 3: Representative image stack. After the imaging experiment the brain was dissected. The neuron from figure 1 is scanned using a confocal microscope with a X20 oil immersion objective, alpha-lobe outlined in purple.

<ol>
	<li>Joerges, J., K&#252;ttner, A., Galizia, C. G., Menzel, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Representations+of+odors+and+odor+mixtures+visualized+in+the+honeybee+brain.">Representations of odors and odor mixtures visualized in the honeybee brain.</a> Nature. 387 (6630), 285-288 (1997).</li><li>Rehder, V. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Quantification+of+the+honeybee's+proboscis+reflex+by+electromyiographic+recordings.">Quantification of the honeybee's proboscis reflex by electromyiographic recordings.</a> J. Insect Physiol. 33, 501-507 (1987).</li><li>Galizia, C. G., Vetter, R. S., Christensen, T. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Optical+Methods+for+Analyzing+Odor-Evoked+Activity+in+the+Insect+Brain.">Optical Methods for Analyzing Odor-Evoked Activity in the Insect Brain.</a> Methods in Insect Sensory Neuroscience. , 345-388 (2004).</li><li>Mauelshagen, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Neural+correlates+of+olfactory+learning+paradigms+in+an+identified+neuron+in+the+honeybee+brain.">Neural correlates of olfactory learning paradigms in an identified neuron in the honeybee brain.</a> J Neurophysiol. 69 (2), 609-625 (1993).</li><li>Kuwabara, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Bildung+des+bedingten+Reflexes+von+Pavlovs+Typus+bei+der+Honigbiene+Apis+mellifica,+Hokaido+Univ.">Bildung des bedingten Reflexes von Pavlovs Typus bei der Honigbiene Apis mellifica, Hokaido Univ.</a> Zool. J. Sci. 13, 458-464 (1957).</li><li>Bitterman, M. E., Menzel, R., Fietz, A., Schafer, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Classical+conditioning+of+proboscis+extension+in+honeybees+(Apis+mellifera.">Classical conditioning of proboscis extension in honeybees (Apis mellifera.</a> Journal of Comparative Psychology. 97 (2), 107-119 (1983).</li></ol>

In this presentation we have gone through all the steps to investigate in vivo calcium signals in the honeybee. We applied this technique to mushroom body neurons, but imaging may be done on all neurons for which a staining technique can be established. We have focused on neurons in the olfactory pathway of the honey bee. The experiments are all performed in a darkroom. The set up is illuminated with a long wave light which lies out of the optical perception range of the bee and does not excite the dyes. It is...

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		<div>
		<table border="1">
		<thead>
		<tr>
		<th>Material Name</th>
		<th>Type</th>
		<th>Company</th>
		<th>Catalogue Number</th>
		<th>Comment</th>
		</tr>
		</thead>
		<tbody>
		
<tr>
<td>low melting point hard wax Deiberit 502</td>
<td>&nbsp;</td>
<td>Dr. B&ouml;hme & Sch&ouml;ps Dental GmbH</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<tr>
<td>FURA-2 dextran potassium salt, 10 000 MW</td>
<td>&nbsp;</td>
<td>Invitrogen/Molecular Probes</td>
<td>F-3029</td>
<td>Protect from light.</td>
</tr>
<tr>
<td>tetramethylrhodamine dextran 10 000 MW</td>
<td>&nbsp;</td>
<td>Invitrogen/Molecular Probes</td>
<td>D-3312</td>
<td>Protect from light.</td>
</tr>
<tr>
<td>n-eicosane</td>
<td>&nbsp;</td>
<td>Sigma-Aldrich</td>
<td>21, 927-4</td>
<td>&nbsp;</td>
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<td>Kwik Sil Adhesive</td>
<td>&nbsp;</td>
<td>WPI</td>
<td>KWIK SIL</td>
<td>&nbsp;</td>
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<td>&nbsp;</td>
<td>TILL Photonics</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
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<td>CCD camera</td>
<td>&nbsp;</td>
<td>Imago</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
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<td>CED</td>
<td>&nbsp;</td>
<td>Texas Instruments</td>
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in vivo ca2+- imaging of mushroom body neurons during olfactory learning in the honey bee

The in vivo and semi-in vivo preparation for Calcium imaging has been developed in our lab by Joerges, K&uuml;ttner and Galizia over ten years ago, to measure odor evoked activity in the antennal lobe1. From then on, it has been continuously refined and applied to different neuropiles in the bee brain. Here, we describe the preparation currently used in the lab to measure activity in mushroom body neurons using a dextran coupled calcium-sensitive dye (Fura-2). We retrogradely stain mushroom body neurons by injecting dye into their axons or soma region. We focus on reducing the invasiveness, to achieve a preparation in which it is still possible to train the bee using PER conditioning. We are able to monitor and quantify the behavioral response by recording electro-myograms from the muscle which controls the PER (M17)2.
After the physiological experiment the imaged structures are investigated in greater detail using confocal scanning microscopy to address the identity of the neurons.

Watch this Scientific Journal Video about In vivo Ca2+- Imaging of Mushroom Body Neurons During Olfactory Learning in the Honey Bee at JoVE.com

In vivo Ca2+- Imaging of Mushroom Body Neurons During Olfactory Learning in the Honey Bee

Bees can be conditioned in an appetitive olfactory learning paradigm (PER-conditioning). Using odors as stimuli, we established a method in which behavior is recorded while simultaneously Calcium Imaging is used to measure odor evoked activity in mushroom body neurons in vivo.

In vivo Ca2+- Imaging of Mushroom Body Neurons During Olfactory Learning in the Honey Bee

The in vivo and semi-in vivo preparation for Calcium imaging has been developed in our lab by Joerges, K&uuml;ttner and Galizia over ten years ago, to ...

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13.9K Views.  Freie Universität Berlin. Le api possono essere condizionate in un paradigma di apprendimento olfattivo appetitivo (PER condizionata). Utilizzando gli odori come stimoli, abbiamo stabilito un metodo in cui il comportamento viene registrato contemporaneamente Imaging calcio è utilizzato per misurare l'attività odore evocato nei neuroni del corpo dei funghi In vivo.

Video: In vivo Ca2+- Imaging of Mushroom Body Neurons During Olfactory Learning in the Honey Bee

Micro-drive Array for Chronic in vivo Recording: Drive Fabrication

Chronic recording of large populations of neurons is a valuable technique for studying the function of neuronal circuits in awake behaving rats. Lightweight recording devices carrying a high density array of tetrodes allow for the simultaneous monitoring of the activity of tens to hundreds of individual neurons. Here we describe a protocol for the fabrication of a micro-drive array with twenty one independently movable micro-drives. This device has been used successfully to record from hippocampal and cortical neurons in our lab. We show how to prepare a custom designed, 3-D printed plastic base that will hold the micro-drives. We demonstrate how to construct the individual micro-drives and how to assemble the complete micro-drive array. Further preparation of the drive array for surgical implantation, such as the fabrication of tetrodes, loading of tetrodes into the drive array and gold-plating, is covered in a subsequent video article.

Micro-drive Array for Chronic in vivo Recording: Drive Fabrication

In this protocol we demonstrate how to fabricate a micro-drive array for chronic electrophysiological recordings in rats.

Chronic recording of large populations of neurons is a valuable technique for studying the function of neuronal circuits in awake behaving rats. ...

Micro-drive Array for Chronic in vivo Recording: Tetrode Assembly

The tetrode, a bundle of four electrodes, has proven to be a valuable tool for the simultaneous recording of multiple neurons in-vivo.  The differential amplitude of action potential signatures over the channels of a tetrode allows for the isolation of single-unit activity from multi-unit signals.  The ability to precisely control the stereotaxic location and depth of the tetrode is critical for studying coordinated neural activity across brain regions.  In combination with a micro-drive array, it is possible to achieve precise placement and stable control of many tetrodes over the course of days to weeks.  In this protocol, we demonstrate how to fabricate and condition tetrodes using basic tools and materials, install the tetrodes into a multi-drive tetrode array for chronic in-vivo recording in the rat, make ground wire connections to the micro-drive array, and attach a protective cone onto the micro-drive array in order to protect the tetrodes from physical contact with the environment.



Micro-drive Array for Chronic in vivo Recording: Tetrode Assembly

In this protocol we demonstrate how to fabricate and condition tetrodes for use with a micro-drive array, which was designed for chronic electrophysiological recordings in rats. In addition, we illustrate the final stages of micro-drive array construction, which includes installing ground wires and a protective cone.

The tetrode, a bundle of four electrodes, has proven to be a valuable tool for the simultaneous recording of multiple neurons in-vivo.  The differential ...

Techniques for Imaging Ca2+ Signaling in Human Sperm

Fluorescence microscopy of cells loaded with fluorescent, Ca2+-sensitive dyes is used for measurement of spatial and temporal aspects of Ca2+ signaling in live cells. Here we describe the method used in our laboratories for loading suspensions of human sperm with Ca2+-reporting dyes and measuring the fluorescence signal during physiological stimulation. Motile cells are isolated by direct swim-up and incubated under capacitating conditions for 0-24 h, depending upon the experiment. The cell-permeant AM (acetoxy methyl ester) ester form of the Ca2+-reporting dye is then added to a cell aliquot and a period of 1 h is allowed for loading of the dye into the cytoplasm. We use visible wavelength dyes to minimize photo-damage to the cells, but this means that ratiometric recording is not possible. Advantages and disadvantages of this approach are discussed. During the loading period cells are introduced into an imaging chamber and allowed to adhere to a poly-D-lysine coated coverslip. At the end of the loading period excess dye and loose cells are removed by connection of the chamber to the perfusion apparatus. The chamber is perfused continuously, stimuli and modified salines are then added to the perfusion header. Experiments are recorded by time-lapse acquisition of fluorescence images and analyzed in detail offline, by manually drawing regions of interest. Data are normalized to pre-stimulus levels such that, for each cell (or part of a cell), a graph showing the Ca2+ response as % change in fluorescence is obtained.

Techniques for Imaging Ca2+ Signaling in Human Sperm

Stimulus-evoked [Ca2+]i signals of individual human sperm are assessed. Motile cells are loaded with Ca2+-sensitive fluorescent dye (AM-ester method) and immobilised in a perfusable chamber. Cells are imaged by time-lapse fluorescence microscopy and stimulated via the perfusing medium. Responses of single cells (or regions) are analysed offline using Excel.

Fluorescence microscopy of cells loaded with fluorescent, Ca2+-sensitive dyes is used for measurement of spatial and temporal aspects of Ca2+ signaling in ...

Rapid Genetic Analysis of Epithelial-Mesenchymal Signaling During Hair Regeneration

Hair follicle morphogenesis, a complex process requiring interaction between epithelia-derived keratinocytes and the underlying mesenchyme, is an attractive model system to study organ development and tissue-specific signaling. Although hair follicle development is genetically tractable, fast and reproducible analysis of factors essential for this process remains a challenge. Here we describe a procedure to generate targeted overexpression or shRNA-mediated knockdown of factors using lentivirus in a tissue-specific manner. Using a modified version of a hair regeneration model 5, 6, 11, we can achieve robust gain- or loss-of-function analysis in primary mouse keratinocytes or dermal cells to facilitate study of epithelial-mesenchymal signaling pathways that lead to hair follicle morphogenesis. We describe how to isolate fresh primary mouse keratinocytes and dermal cells, which contain dermal papilla cells and their precursors, deliver lentivirus containing either shRNA or cDNA to one of the cell populations, and combine the cells to generate fully formed hair follicles on the backs of nude mice. This approach allows analysis of tissue-specific factors required to generate hair follicles within three weeks and provides a fast and convenient companion to existing genetic models.

Tissue-specific analysis of a hair follicle regeneration model using lentivirus to mediate gain- or loss-of-function.

Hair follicle morphogenesis, a complex process requiring interaction between epithelia-derived keratinocytes and the underlying mesenchyme, is an ...

Live Imaging of Apoptotic Cell Clearance during Drosophila Embryogenesis

The proper elimination of unwanted or aberrant cells through apoptosis and subsequent phagocytosis (apoptotic cell clearance) is crucial for normal development in all metazoan organisms. Apoptotic cell clearance is a highly dynamic process intimately associated with cell death; unengulfed apoptotic cells are barely seen in vivo under normal conditions. In order to understand the different steps of apoptotic cell clearance and to compare 'professional' phagocytes - macrophages and dendritic cells to 'non-professional' - tissue-resident neighboring cells, in vivo live imaging of the process is extremely valuable. Here we describe a protocol for studying apoptotic cell clearance in live Drosophila embryos. To follow the dynamics of different steps in phagocytosis we use specific markers for apoptotic cells and phagocytes. In addition, we can monitor two phagocyte systems in parallel: 'professional' macrophages and 'semi-professional' glia in the developing central nervous system (CNS). The method described here employs the Drosophila embryo as an excellent model for real time studies of apoptotic cell clearance.

Live Imaging of Apoptotic Cell Clearance during Drosophila Embryogenesis

Here we describe an effective method for studying dynamics of apoptotic cell clearance in vivo. This method employs live Drosophila embryos as a powerful model for monitoring phagocytosis of apoptotic cells using specific labeling of apoptotic cells and phagocytes.

The  proper elimination of unwanted or aberrant cells through apoptosis and  subsequent phagocytosis (apoptotic cell clearance) is crucial for normal  ...

Live Imaging Assay for Assessing the Roles of Ca2+ and Sphingomyelinase in the Repair of Pore-forming Toxin Wounds

Plasma membrane injury is a frequent event, and wounds have to be rapidly repaired to ensure cellular survival. Influx of Ca2+ is a key signaling event that triggers the repair of mechanical wounds on the plasma membrane within ~30 sec. Recent studies revealed that mammalian cells also reseal their plasma membrane after permeabilization with pore forming toxins in a Ca2+-dependent process that involves exocytosis of the lysosomal enzyme acid sphingomyelinase followed by pore endocytosis. Here, we describe the methodology used to demonstrate that the resealing of cells permeabilized by the toxin streptolysin O is also rapid and dependent on Ca2+ influx. The assay design allows synchronization of the injury event and a precise kinetic measurement of the ability of cells to restore plasma membrane integrity by imaging and quantifying the extent by which the liphophilic dye FM1-43 reaches intracellular membranes. This live assay also allows a sensitive assessment of the ability of exogenously added soluble factors such as sphingomyelinase to inhibit FM1-43 influx, reflecting the ability of cells to repair their plasma membrane. This assay allowed us to show for the first time that sphingomyelinase acts downstream of Ca2+-dependent exocytosis, since extracellular addition of the enzyme promotes resealing of cells permeabilized in the absence of Ca2+.

Live Imaging Assay for Assessing the Roles of Ca2+ and Sphingomyelinase in the Repair of Pore-forming Toxin Wounds

Live imaging of cells exposed to the lipophilic dye FM1-43 allows precise determination of the kinetics by which pore-forming toxins are removed from the plasma membrane. This is a sensitive assay that can be used to assess requirements for Ca2+, sphingomyelinase and other factors on plasma membrane repair.

Plasma membrane injury is a frequent event, and wounds have to be rapidly repaired to ensure cellular survival. Influx of Ca2+ is a key signaling event ...

Obtaining Specimens with Slowed, Accelerated and Reversed Aging in the Honey Bee Model

Societies of highly social animals feature vast lifespan differences between closely related individuals. Among social insects, the honey bee is the best established model to study how plasticity in lifespan and aging is explained by social factors.
The worker caste of honey bees includes nurse bees, which tend the brood, and forager bees, which collect nectar and pollen. Previous work has shown that brain functions and flight performance senesce more rapidly in foragers than in nurses. However, brain functions can recover, when foragers revert back to nursing tasks. Such patterns of accelerated and reversed functional senescence are linked to changed metabolic resource levels, to alterations in protein abundance and to immune function. Vitellogenin, a yolk protein with adapted functions in hormonal control and cellular defense, may serve as a major regulatory element in a network that controls the different aging dynamics in workers.
Here we describe how the emergence of nurses and foragers can be monitored, and manipulated, including the reversal from typically short-lived foragers into longer-lived nurses. Our representative results show how individuals with similar chronological age differentiate into foragers and nurse bees under experimental conditions. We exemplify how behavioral reversal from foragers back to nurses can be validated. Last, we show how different cellular senescence can be assessed by measuring the accumulation of lipofuscin, a universal biomarker of senescence.
For studying mechanisms that may link social influences and aging plasticity, this protocol provides a standardized tool set to acquire relevant sample material, and to improve data comparability among future studies.

In honey bee workers, aging depends on social behaviors rather than on chronological age. Here we show how worker-types with very different aging patterns can be obtained and analyzed for cellular senescence.

Societies of highly social animals feature vast lifespan differences between closely related individuals. Among social insects, the honey bee is the best ...

Methods for Comparing Nutrients in Beebread Made by Africanized and European Honey Bees and the Effects on Hemolymph Protein Titers

Honey bees obtain nutrients from pollen they collect and store in the hive as beebread. We developed methods to control the pollen source that bees collect and convert to beebread by placing colonies in a specially constructed enclosed flight area. Methods were developed to analyze the protein and amino acid composition of the pollen and beebread. We also describe how consumption of the beebread was measured and methods used to determine adult worker bee hemolymph protein titers after feeding on beebread for 4, 7 and 11 days after emergence. Methods were applied to determine if genotype affects the conversion of pollen to beebread and the rate that bees consume and acquire protein from it. Two subspecies (European and Africanized honey bees; EHB and AHB respectively) were provided with the same pollen source. Based on the developed methods, beebread made by both subspecies had lower protein concentrations and pH values than the pollen. In general, amino acid concentrations in beebread made by either EHB or AHB were similar and occurred at higher levels in beebread than in pollen. Both AHB and EHB consumed significantly more of the beebread made by AHB than by EHB. Though EHB and AHB consumed similar amounts of each type of beebread, hemolymph protein concentrations in AHB were higher than in EHB. Differences in protein acquisition between AHB and EHB might reflect environmental adaptations related to the geographic region where each subspecies evolved. These differences could contribute to the successful establishment of AHB populations in the New World because of the effects on brood rearing and colony growth.

Here are methods to quantify nutrients in pollen before and after its conversion to beebread by two subspecies of honeybees. We describe techniques to measure beebread consumption and resulting protein titers in both subspecies.

Honey bees obtain nutrients from pollen they collect and store in the hive as beebread. We developed methods to control the pollen source that bees ...

Imaging Local Ca2+ Signals in Cultured Mammalian Cells

Cytosolic Ca2+ ions regulate numerous aspects of cellular activity in almost all cell types, controlling processes as wide-ranging as gene transcription, electrical excitability and cell proliferation. The diversity and specificity of Ca2+ signaling derives from mechanisms by which Ca2+ signals are generated to act over different time and spatial scales, ranging from cell-wide oscillations and waves occurring over the periods of minutes to local transient Ca2+ microdomains (Ca2+ puffs) lasting milliseconds. Recent advances in electron multiplied CCD (EMCCD) cameras now allow for imaging of local Ca2+ signals with a 128 x 128 pixel spatial resolution at rates of &#62;500 frames sec-1 (fps). This approach is highly parallel and enables the simultaneous monitoring of hundreds of channels or puff sites in a single experiment. However, the vast amounts of data generated (ca. 1 Gb per min) render visual identification and analysis of local Ca2+ events impracticable. Here we describe and demonstrate the procedures for the acquisition, detection, and analysis of local IP3-mediated Ca2+ signals in intact mammalian cells loaded with Ca2+ indicators using both wide-field epi-fluorescence (WF) and total internal reflection fluorescence (TIRF) microscopy. Furthermore, we describe an algorithm developed within the open-source software environment Python that automates the identification and analysis of these local Ca2+ signals. The algorithm localizes sites of Ca2+ release with sub-pixel resolution; allows user review of data; and outputs time sequences of fluorescence ratio signals together with amplitude and kinetic data in an Excel-compatible table.

Imaging Local Ca2+ Signals in Cultured Mammalian Cells

Here we present techniques for imaging local IP3-mediated Ca2+ events using fluorescence microscopy in intact mammalian cells loaded with Ca2+ indicators together with an algorithm that automates identification and analysis of these events.

Cytosolic Ca2+ ions regulate numerous aspects of cellular activity in almost all cell types, controlling processes as wide-ranging as gene transcription, ...

High-resolution Time-lapse Imaging and Automated Analysis of Microtubule Dynamics in Living Human Umbilical Vein Endothelial Cells

The physiological process by which new vasculature forms from existing vasculature requires specific signaling events that trigger morphological changes within individual endothelial cells (ECs). These processes are critical for homeostatic maintenance such as wound healing, and are also crucial in promoting tumor growth and metastasis. EC morphology is defined by the organization of the cytoskeleton, a tightly regulated system of actin and microtubule (MT) dynamics that is known to control EC branching, polarity and directional migration, essential components of angiogenesis. To study MT dynamics, we used high-resolution fluorescence microscopy coupled with computational image analysis of fluorescently-labeled MT plus-ends to investigate MT growth dynamics and the regulation of EC branching morphology and directional migration. Time-lapse imaging of living Human Umbilical Vein Endothelial Cells (HUVECs) was performed following transfection with fluorescently-labeled MT End Binding protein 3 (EB3) and Mitotic Centromere Associated Kinesin (MCAK)-specific cDNA constructs to evaluate effects on MT dynamics. PlusTipTracker software was used to track EB3-labeled MT plus ends in order to measure MT growth speeds and MT growth lifetimes in time-lapse images. This methodology allows for the study of MT dynamics and the identification of how localized regulation of MT dynamics within sub-cellular regions contributes to the angiogenic processes of EC branching and migration.

Protocols for Human Umbilical Vein Endothelial Cell (HUVEC) culture, transient transfection of fluorescently-labeled markers of microtubule growth, live-cell imaging and automated analysis of interphase microtubule growth dynamics are detailed.

The physiological process by which new vasculature forms from existing vasculature requires specific signaling events that trigger morphological changes ...