Lab Protocols

This protocol explains how to: use Olympus fluorescent microscopes to image in vitro experiment slides.   Protocol written by: Julie Ruble Turn on green burner power button first, and then the microscope and then the camera. Start viewing in fluorescence (change filters using the filter wheel until you reach the green filter). Begin at the corner of the slide. Use the 40X objective lens. When looking for neurons to image, scan through the slide methodically so as not to miss any: When you find a neuron, position it with the cell body in the field of vision but off to the side (so you'll be able to get more neurite in the picture and therefore take fewer pictures). NOTE: When you are not looking at or imaging the neuron, close the shutter on the right side of the microscope so as not to bleach the neuron. To take a picture, open SPOT (the program may have a shortcut on the desktop, but you may need to find it in the C folder and create one). Then, to take a flu

How to clean an objective:  Turn off the microscope. Turn the nosepiece until the objective in question is at an angle proper for removal or cleaning (See Figure 1).     Figure 1. If you choose to remove the objective for cleaning, very carefully unscrew it from the nosepiece and gently place it on a Kim-wipe on a clean, flat surface. Whether the objective is still attached to the nosepiece or not, the next step is to cover a Q-tip in lens paper and soak the tip in either alcohol or lens cleaning solution. (Lens-cleaning solution can be found in a small, zippered bag on the shelf for the Olympus microscope in Dana) (See Figures 2 and 3). Figure 2.   Figure 3 . Gently rub the glass on the objective—the part that is adjacent to the sample when viewing- with the wet Q-tip. Do NOT touch the inside of the objective where it attaches to the microscope. When you feel that you have sufficiently cleaned the dirt or oil from the objective, acquire lens-cleaning paper It can be found in

This protocol explains how to: add a scalebar to an image. Protocol written by: Barbara Lom 1. To include a scale bar on any image you need to acquire the image and an image of the micrometer (tiny ruler) slide at the same magnification and image parameters. 2. Open your "ruler" image in Photoshop. If the image is a bit askew you will need to rotate the image so that the ruler marks are "square" in the frame of your picture (it is OK to crop the ruler image if necessary in this step). 3. Go to the "layers" box (usually at the lower right - if not then make layers visible under the "window" menu & select "show layers") 4. Click on the arrow in the circle in the upper right corner of the layers box and select "new layer" 5. Go to the toolbar and select the pencil tool (which is sometimes hiding under the paintbrush tool). 6. Click on brush arrow (above the image) to change the "width"

This protocol explains how to: align the phase on the Olympus fluorescent microscope.   Protocol written by: Barbara Lom, Fiona Watson, Julie Ruble NOTE: While following this protocol, please reference the labeled diagram found below.  A full-sized version of this image is available here . Start with the 10X objective and make sure the Phase Selector Ring is on "Ph 1."  Focus on a slide. Take out the right eyepiece of the Olympus fluorescent microscope and insert the UT30 centering telescope (found bubble-wrapped in a plastic jar by one of the scopes). Push in and turn allen wrench knobs until both rings viewed through the telescope eyepiece align in the dark circle.  Make sure the allen wrenches are catching in the screw hole when you push them in to turn them. Repeat this with each objective and its corresponding Phase Selector Ring setting.  For the 40X objective, the ring should be on "Ph 2," and for a 100X objective, it should be set on "Ph 3.

This protocol explains how to: take pictures with the Nikon inverted scopes using ImagePro.   Protocol written by: Barbara Lom Turn on the power to the microscope and check that the light is directed to the eyepieces by setting the dial on the upper right hand side to the “bino” setting. Use the 10X objective to find and focus on your sample.  Adjust the light intensity is adjusted with the dial on the lower/front/left of the microscope.  If you are using phase contrast optics make sure the phase ring is in place (slider near the top of the microscope). Adjust the focus and lighting to get your best phase contrast image. Divert light from oculars to the camera by turning the dial from “bino” to “photo” setting. Turn the camera power on (small beige box). Turn on the PC and log into Windows. Open ImagePro+ (Microscope Icon – upper left side of desktop). Select the “complete” option and click “OK”.  Ignore the registration request and close the macro player window if

This protocol explains how to: Image Using a 100x Objective (Oil Immersion) Protocol revised: 11/17/10 Protocol written by: Aaron Deal Note: When imaging a slide using a 100x objective, oil is required for a precise image. 1. View the slide on a compound microscope. Begin on a lower powered objective and then move up to a dry 100x objective in order to find the desired, or potentially desired, image. (If the image is brightfield, see the Kohler illumination protocol for instructions on setting up Kohler illumination) 2. Once the image is in view, rotate the nosepiece so that no objective is pointing towards the slide. 3. Place 1-2 drops of objective lens oil (located on the shelves above the microscope) directly onto the slide where the objective will be located. 4. Rotate the nosepiece back to the 100x objective, without passing any other objectives through the oil (the 100x objective lens should now be surrounded by the oil). 5. The stage can still be moved, but a reappli

Use this protocol if you need to prepare a specimen for imaging with a microscope that has a small working distance (such as the confocal) and preserve it for storage afterward. 1) Gather the supplies you'll need: (also listed below this image) Microscope slides - preferably from a box of plain, precleaned, slides with the dimensions 25x75x1mm Plastic pipette (or glass, if you prefer) Sharpie or another permanent marker Nail polish Forceps (with black tape on the handle, which designates use with fixed specimens only) Reinforcement labels Slide covers (coverslips) - preferably from a box that lists the dimensions as "22x22-1.5um" Fluoro-gel 2) Obtain your prepared (fixed, sliced) samp le. 3) On a slide, label your initials, experiment, and date with your permanent marker: 4) Place one or two reinforcement labels on the slide. If you choose only one, place it in the center of the slide; if two, place them evenly spaced out. The latter is usually preferred

1) Open Photoshop on the computer 2) Click File > Open 3) Select the images that you want to overlay and open them in photoshop 4) Make sure there is a saved copy of all the original images 5) Save the images under new names before you begin manipulating them, so that you will always have an unchanged original 6) Once you have opened all of your images, pick one to be your base image (In this example, the base image will be green, and the other two images will be red and blue). 7) Bring the blue fluoresent image to the front 8) Go to Image> Select> All 9) Then, go to File> Copy (or Ctrl C) 10) Once you have a copy of the blue image, go back to your base green image and click on the channels tab (next to the layers tab at the bottom right) 11) Highlight the blue channel 12) Paste (File> Paste or Ctrl V) your copied blue image into the blue channel 13) Repeat these steps copying the red image into the red channel 14) Now save the overlay image as a photoshop a

1) Open Photoshop on the computer 2) Click File > Open 3) Select the images that you want to overlay and open them in photoshop 4) Make sure there is a saved copy of all the original images 5) Save the images under new names before you begin manipulating them, so that you will always have an unchanged original 6) Once you have opened all of your images, decide how many panels you need for your final image, and pick one to be your base image 7) On your base image, go to Image> Canvas Size 8) A new box will open with the height and width of your current canvas 9) Multiply the height or width by the number of images that need to fit within your canvas (This means if you want a three panel image multiply the width by three and do not change the height. If you want a four panel square image, multiply the width and height by two). 10) Once you change the canvas height and width to the desired size, anchor the base image where you want it in the final image panel (Ex. If you ar

100% Steinberg’s Solution (AKA 5xMR) (makes 2 liters) 40 ml NaCl stock solution (17% w/v NaCl) 20 ml KCl stock solution (0.5% w/v KCl) 20 ml CaCl2 stock solution (0.5% w/v CaCl2) 20 ml MgSO4 stock solution (1.025% w/v MgSO4) 1 ml gentamycin stock solution (4% w/v gentamycin) 2.4 g HEPES 1899 ml ddH2O Adjust pH to 7.4 - 7.6 and store in refrigerator. Usually, you will make 2-4 liters at a time. To make the required stock solutions,

Treat tads with ~1.5 ml PTU prior to stage 30-32 to prevent pigmentation. 0.3% PTU 0.3 g Phenylthiocarbamide (Sigma # P-7629, also called Phenylthiourea) 100 ml 20% Steinberg’s solution Some of the PTU will not dissolve. Shake solution prior to use to dissolve some of the precipitate (put “shake well” on label). Store in refrigerator.

100 ml 10X MBS stock 1.25 ml 4% gentamycin (1 g gentamycin sulfate in 25 ml ddH2O) 893 ml ddH2O 7 ml autoclaved 0.1M CaCl2 (11.1 g CaCl2 / 100 ml ddH2O) Adjust pH to ~7.4. Sterile filter prior to use. Make fresh weekly and store in refrigerator.

Modified Barth’s Saline Operating Solution (MBS) (makes 1 liter) 100 ml 10X MBS stock 500 ul 1% phenol red (pH indicator, if needed) 10 ml pen/strep antibiotic cocktail 5 ml 4% gentamycin (1 g gentamycin sulfate in 25 ml ddH2O) 0.2 g Ethyl 3-aminobenzoate methanesulfonic acid salt (C9H11NO2*CH4SO3) 7 ml autoclaved 0.1M CaCl2 (11.1 g CaCl2 / 100 ml ddH2O) Fill to 1000 ml with ddH2O. Adjust pH to ~7.4. Sterile filter immediately prior to use. Make fresh each week and store in refrigerator. NOTE: the anesthesia amount may need to be decreased for some tads.

2% Agar (for agar dishes) 2 g Agar 100 ml ddH2O Microwave to dissolve the agar, lightly swirling the mixture periodically. Turn off the room light to see the solution better; it will boil over if not watched carefully. Stop the microwave when the solution starts to boil up in the flask and swirl gently. Be careful while swirling the solution as it may unexpectedly boil over due to the added kinetic energy. Use orange heat resistant gloves to handle the flask. Once the solution has cooled a bit, microwave again, stopping when it boils. Repeat this several times – the more boiling, the less bubbles in your finished agar dishes. Pour the hot mixture into petri dish lids and let it set. Store the dishes in an air tight container in the fridge. Make new every two weeks.

1X Tadpole Anesthesia (0.05%) 10 ml 10X Tadpole Anesthetic 90 ml 20% Steinberg’s solution or 0.05g Ethyl 3-aminobenzoate methanesulfonic acid salt (C9H11NO2*CH4SO3) 100 ml 20% Steinberg’s solution Adjust pH to ~7.4.

use this anesthesia to anesthetize adult frogs before dissecting out testes, and also for 1X dilution. 10X Tadpole Anesthesia (0.5%) (AKA Adult Anesthetic, MS222, Tricane) 0.5 g Ethyl 3-aminobenzoate methanesulfonic acid salt (C9H11NO2*CH4SO3) 100 ml 20% Steinberg’s solution Initially the solution is very acidic. Be careful adjusting the pH, as it tends to jump quickly past pH 4.5. Our meter has trouble reading anesthesia and will never truly stabilize. Once the “S” symbol appears on meter, use pH paper to adjust pH to ~7.4.

We use this to anesthetize tadpoles before injections, etc. 1X Tadpole Anesthesia (0.05%) 10 ml 10X Tadpole Anesthetic 90 ml 20% Steinberg’s solution or 0.05g Ethyl 3-aminobenzoate methanesulfonic acid salt (C9H11NO2*CH4SO3) 100 ml 20% Steinberg’s solution Adjust pH to ~7.4.

Culture Media (makes 100 ml) 10 ml Fetal Bovine Serum (FBS) 1 ml pen/strep antibiotic cocktail (AKA Fungibact) 1 ml Embryo Extract solution (EE) 28 ml ddH2O 60 ml L15 Adjust pH to ~7.4. Sterile Filter prior to using; do not autoclave.

Dilute to 1X to yield zebrafish embryo rearing solution. This embryo medium contains antibiotics for fish that are susceptible to diseases (like transgenic and fluorescent fish). For the standard embryo medium without antibiotics, see 60X E3 stock. 20X E2 stock (makes 2 liters) 35 g NaCl 1.5 g KCl 0.82 g KH2PO4 4.8 g MgSO4 0.24 Na2HPO4 2000 ml ddH2O Autoclave (optional) and store in the refrigerator. Make the following 3 solutions. When diluting to 1X E2, combine 50 ml 20X E2, 2 ml CaCl2 solution, 2 ml of the NaHCO3 solution, and ddH2O to 1 liter. Add antibiotic diluted 1:500 in1X E2. 1. Add 14.5 g CaCl2 in 200 ml ddH2O, autoclave, and refrigerate. 2. Add 6 g NaHCO3 to 200 ml ddH2O, autoclave, and refrigerate. 3. Make a 60 mg/ml penicillin, 100 mg/ml streptomycin stock, aliquot and store at -20o C. Dilute 1:500 in 1X E2 for use.

Serum Free Culture Media (makes 100 ml) 1 ml pen/strep antibiotic cocktail (AKA Fungibact) 0.1 g Albumin, bovine serum (fatty acid free, Sigma# A8806) 39 ml ddH2O 60 ml L15 Adjust pH to ~7.4. Sterile Filter prior to using; do not autoclave.

Reaggregation Solution with 0.05% Anesthesia (makes 1 liter) 0.56 g Tris Base 0.32 g MgSO4 (7H2O) 6.81 g NaCl 0.05 g KCl 1.47 g CaCl2 (dihydrate) 1000 ml ddH2O 0.5 g Ethyl 3-aminobenzoate methanesulfonic acid salt (C9H11NO2*CH4SO3) Phenol Red (a few drops to indicate pH) Adjust pH to ~7.4. Sterile Filter prior to using; do not autoclave.

Low [Ca++] Reaggregation Solution with 0.05% Anesthesia (makes 1 liter) 6.81 g NaCl 0.05 g KCl 1.47 g CaCl2 (dihydrate) 1.91 g HEPES 1000 ml ddH2O 0.5 g Ethyl 3-aminobenzoate methanesulfonic acid salt (C9H11NO2*CH4SO3) Phenol Red (a few drops to indicate pH) Adjust pH to ~7.4. Sterile Filter prior to using; do not autoclave.

Disaggregation Solution (makes 1 liter) 6.81 g NaCl 0.05 g KCl 0.56 g Tris Base 0.117 g EDTA 1000 mL ddH2O Phenol Red (a few drops to indicate pH) Adjust pH to ~7.4. Sterile Filter prior to using; do not autoclave.

Embryo Extract Solution (EE) (makes 100 ml) 60 % L15 10 % FBS (Fetal Bovine Serum) 30 % ddH2O 1 % FB (Fungibact; Antibiotic antimycotic) 1. Remove jelly coat and membrane from embryos (stage 23-30) and count (you will need the number of embryos later). 2. Place embryos in tissue grinder, drawing off as much liquid as possible. 3. Add ~1 squirt of EE media. 4. Homogenize embryos (~ 50 plunges). Push down slowly and pull up quickly. 5. Transfer to 2 centrifuge tube and increase volume to ~ 30 mls using EE media or about 1/2 full. Balance both tubes. 6. Spin at 15K, 4C for 90 mins in SS-34 rotor. 7. Three layers appear. Bottom = Crude, Middle = Liquid, Top= Milky lipid layer. Carefully pipette middle layer. Avoid the top layer because it clogs the filter. 8. Add EE media such that the final concentration is 1 ml / 1 embryo. 9. Sterile filter.

PBT 100 ml 1X PBS g BSA (bovine serum albumin Cohn Analog (Sigma# A1470)) 100 ul triton Make fresh each use. Can be stored in refrigerator for no more than 24 hours.

0.4 M PO4 Buffer Solution (makes 500 ml) 5.25 g NaH2PO4 * H2O 43.3 g Na2HPO4 * 7H2O (or 22.94 g Na2HPO4) 500 ml ddH2O Adjust pH to ~ 7.4.

0.1 M PO4 Buffer Solution (makes 400 ml) 100 ml 0.4 M PO4 buffer solution 300 ml ddH2O Adjust pH to ~ 7.4.

This solution is used in AChE stain solution. Acetate Buffer (makes 200 ml) 1.353 g NaC2H3O2 (or 0.82 g / 100 ml ddH2O) 0.1995 ml glacial acetic acid (or 0.57 ml / 100 ml ddH2O) 200 ml ddH2O (or 165 ml of sodium solution and 35 ml of acid solution) Adjust pH to 5.3.

This solution is used in AChE stain solution. Glycine/copper solution (makes 100 ml) 3.75 g C2H5NO2 (Glycine) 1.6g CuSO4 100 ml ddH2O

This solution is used in AChE stain solution. 8.8 mM Promethazine (aka phenegram) 50 mg promethazine = 2 crushed tablets 17.8 mL ddH2O Must be made fresh weekly.

his solution is used in AChE stain solution. 0.1 M Copper sulfate (makes 100 ml) 1.5 g CuSO4 100 ml ddH2O

AChE stain solution (makes ~98 ml) AChE Solution 1 (will be blue) 40 ml 0.1 M acetate buffer 28 mL ddH2O 3.2 mL glycine/copper solution 2 ml 8.8 mM promethazine AChE Solution 2 (will be brown/yellow) 18 ml ddH2O 372 mg acetylthiocoline iodide 6 ml 0.1 M copper sulfate Both solutions must be made fresh each time. Filter solution 2 with filter paper and then mix with solution 1. Cover tissue/slides in the combined solution and place on rocker/belly dancer for 24-72 hours, then rinse tissue/slides in ddH2O for five minutes. Repeat rinse three times.

2% Sodium sulfide solution (makes 200 ml) 4 g Na2S * 9H2O 200 ml ddH2O

This solution is used to remove the jelly coat from fertilized Xenopus laevis eggs, allowing you to separate out the good eggs. Cysteine Solution (makes enough to cysteine 1 petri dish of fertilized eggs) 0.6 g Cysteine 1.0 g Tris base 30 ml 20% Steinberg’s solution For 4 dishes, and with our current (2009) basic-leaning water: 1.8g Cysteine 1.7g Tris base 90 ml 20% Steinberg's solution ***Resulting pH: ~7.52

NOTE: This solution should be made fresh before immunostaining. PBDT: 1% DMSO 0.1% Tween-20 Fill with 1X PBS to final volume

Dilute to 1X to yield zebrafish embryo rearing solution. For an embryo medium that contains antibiotics for fish that are susceptible to diseases (like transgenic and fluorescent fish), see 20X E2 stock. 60X E3 stock (makes 2 liters) 34.4 g NaCl 1.52 g KCl 5.8 g CaCl2.2H2O 9.8 g MgSO4.7H2O add 18 MOhm "good" double distilled water up to 2000 ml Store in refrigerator. To dilute to 1X for rearing zebrafish, use 160 ml of stock and fill to 10 liters with ddH2O. This solution is also called "fish juice" by Amy Becton - she will need the lab to make a 2L bottle every week or two for downstairs in fishland more details on this recipe can be found in the Nusslein-Volhard & Dahn purple Zebrafish book page 22 It is helpful to make up about 10 containers of the salts at one time. I just put them in a plastic container with a lid and label them "60X E3 Salts." This way you don't have to worry about weighing them out when Amy needs a new bottl

10 g Tryptone 5 g Yeast Extract 10 g NaCl Bring to 1 liter with Millipore water Adjust pH to 7.5 Pour into multiple small bottles and autoclave with lid loosened (Usually increments of 200-250 mls)

5 g Tryptone 2.5 g Yeast extract 5 g NaCl Bring to 500 mls with Millipore water While mixing, prepare a 1-liter flask with 7.5 g of agar Adjust LB to pH 7.5 with dilute NaOH Pour liquid into flask with agar Cover with aluminum foil and secure with autoclave tape Autoclave 30 min Allow to cool with slow mixing to about 50C (should be able to leave your hands on the glass without it hurting) Meanwhile, remove 1 sleeve of plates from the plastic and save the plastic Add antibiotic and mix again for a few minutes (Ampicillin at 100 ug/ml, Kanamycin at 50 ug/ml, or Chloramphenicol at 25 ug/ml) Pour into agar plates, pouring just a little more than it takes to cover the bottom of the plate Leave at room temperature overnight to reduce condensation Store inverted in platic sleeves at 4C (refrigerator)

BT-Fix Warning: Do not make BT-Fix alone the first time. 0.26 g NaH2PO4 * H2O 1.14 g Na2HPO4 100 ul of 0.12M CaCl2 (1.3 g CaCl2 / 100 ml ddH2O or 1.76 g CaCl2-H2O/100 mL ddH2O) 4 g Sucrose 4 g Paraformaldehyde (Extremely Toxic – Hood, Gloves, Mask, Coat required) This recipe is for 100 mL of BT fix. Put all ingredients except paraformaldehyde into a 100 ml graduated cylinder (if you are doubling the recipe, you need a graduated cylinder that goes up to at least 200 ml). The 0.12M CaCl2 can be mixed in advance and kept in the refrigerator. Fill the graduated cylinder to almost 100 ml with ddH2O (you need to leave room for the rest of the ingredients, so don't fill it all the way to 100 ml. You can always add more later). Place the stirplate under the hood with a fix beaker on it. Place a fix thermometer into the fix beaker. Pour the mixture you just made into the fix beaker. Put a fix stir bar into the beaker and turn on the stirplate. Turn the heat on to about

(makes 400 ml) 100 ml 0.4 M PO4 buffer solution 300 ml ddH2O Adjust pH to ~ 7.4

Tricaine Solution: 400 mg tricaine in 97.9 ml E3 0.26 g Tris in 2.1 ml E3 Adjust pH to ~7 Store this solution in a 50 ml conical tube in the freezer To use as an anesthetic : 4.2 ml tricaine solution ~100 ml E3

20X SSC: Makes 1 Liter . Start with 800 mLs of Millipore H2O Add: 175.3 g NaCl and 88.2 g sodium citrate Dissolve well pH to 7.0 Bring volume to 1 Liter

You will need: Bucket of ice 100ng-500ng (~2 uL) of plasmid DNA to be transformed in a sterile eppendorf tube Competent bacteria cells (DO NOT THAW until ready to begin) Heat block set to 42C LB Broth Bunsen Burner Micropipettors and pipette tips 37C water bath 2 LB agar plates with appropriate antibiotic 37C plate incubator 1. Place tube with plasmid DNA and a tube of competent cells on ice. 2. Allow competent cells to thaw on ice. Do not use your hands to speed it up. 3. As soon as they are thawed, add 50-100uL of the cells to the tube containing the DNA. 4. Incubate on ice for 15'. 5. Heat-shock the cells by placing in the 42C heat block for 90 seconds. 6. Return the tube to ice. 7. Incubate on ice for 2'. 8. Add 300uL of sterile LB broth to the tube using sterile technique. 9. Incubate in a 37C water bath for 45'-1 hour. 10. Plate an aliquot of the cells on LB/antibiotic plates. 11. Invert plates and grow at 37C overnight (Not more than 18-20 hour

Beginning Locate the recipe of the solution you need by scanning the " Recipes " category of the protocol website or using the " Search " tab. Collect all the reagents you need to make your solution.  Room temperature reagents will usually be located in the chemical cabinet (shown below) in alphabetical order while cold or frozen reagents may be kept in the lab fridge or small freezer, respectively. Figure 1. Chemical cabinet. Evaluate the amount of solution you should make.  For solutions used in large amounts (e.g. 20% Steinberg's, 1X PBS) it may make sense to mix up several liters at a time.  For solutions that degrade quickly (e.g. Testes Steinberg's, cysteine) or are used less frequently (e.g. disag, 0.1M PB) you may decide to mix up a smaller amount. If you are making a different volume of solution than the recipe specifies, do not trust yourself to make the calculations and convert the recipe as you go.  Calculate how much of each reagent

0.1 M TEA: TEA = Triethanolamine You will dilute TEA 1:10 in Millipore water: Pour 50 mL TEA into a 50 mL conical tube (TEA is very viscuous, so best to measure it in something that is disposable). Pour TEA from conical tube into 450 mL Millipore water with stirring. pH to 8.0

Location St. Mary's Medical Center, San Francisco, CA Position Type Part-time Position Level Staff Shift: 8HOUR Hours per Pay Period: 8 Facility: ST MARYS MEDICAL CENTER Facility Information: St. Mary's Medical Center is a full-service acute care facility with more than 575 physicians and 1,100 employees who provide high-quality and affordable health care services to the Bay Area community. Home to advanced medical practices, such as the nation's first digital cardiac catheterization laboratory, pioneering spine surgery and comprehensive rehabilitation, St. Mary's Medical Center is one of San Francisco's leading hospitals, offering patients a full range of outpatient and inpatient services delivered with the human touch. Strategies and business development are centered around Oncology Services, Cardiac Services and Orthopedics. Position Requirements: JOB DUTIES: Performs chemical, microscopic,immunologic and microbiological procedures. Evaluates the

Location HCA West Florida Division, Palm Harbor, FL Position Type Full-time Primary Location : United States-Florida-Brooksville Description Oak Hill Hospital, part of the HCA West and Central Florida hospital system, is a 214-acute care hospital in the community of Brooksville. Our experienced healthcare team is dedicated to diagnosing and treating patients faster than any hospital in the county. The hospital is located near the family-friendly town of Brooksville, Oak Hill Hospital is situated on Florida's Nature Coast, which offers fishing, boating, bird watching and hiking. Join us in the Laboratory as a Medical Technologist. The Medical Technologist performs laboratory testing in one or more sections of the laboratory that assists with the diagnosis of patients. The tasks and responsibilities include: * Processes specimens, prepares reagents, performs testing procedures; reports and interprets test results. * Performs quality control testing, instrument ma

Job Title Lab Equipment & Metrology Technician Posted 27/05/2011 (14:21) Agency/Employer AssimilateLtd Description The Lab Equipment & Metrology Technician is responsible for the maintenance and upkeep of a wide range of laboratory equipment within the Biotechnology and CMC-PDS divisions of this global contract research services company. The candidate has a responsibility to conduct all work in compliance with applicable regulatory requirements, and specifically those regulations (GxP) specified in the study plans, protocols or work agreements governing the work in which they are involved. Key Responsibilities Reporting to GMP Equipment & Metrology Manager: Perform routine servicing and calibration of a wide range of equipment; To ensure all equipment is cali

Validation Engineer 05-26-2011 California-Oakland/East Bay Pharmaceutical Assistant QA Supervisor-2nd Shift 05-26-2011 Pennsylvania-Pittsburgh Food & Beverage GC and Analytical Chemists 05-26-2011 Pennsylvania-Philadelphia Multiple Categories Biotechnology Chemical Industry Medical Device Laboratory Technician 05-26-2011 Delaware-Delaware Multiple Categories Medical Device Pharmaceutical Laboratory Technician 05-26-2011 New Jersey-Southern Multiple Categories Biotechnology Chemical Industry Food & Beverage Materials Science Personal Care Pharmaceutical Regulatory Affairs Director 05-25-2011 California-San Diego Medical Device Regulatory Affairs Manager 05-25-2011 California-San Diego Medical Device