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Materials

1. Glass culture tubes with metal caps and labels

• Growth medium, from media room or customized
• Glass pipette tubes
• Parafilm

Equipment

1. Vortexer

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METHOD

1. Place 2 mL of the appropriate sterile medium in a 13 mm yellow-capped culture tube. If more culture is needed, place up to 5 mL in a 16 mm green-capped culture tube. Large amount of culture should be made in growth flasks, using up to 25% flask volume in medium (e.g., no more than 30 mL in a 125 mL flask, or 250 mL in a 1000 mL flask).

• Touch a single colony with the end of an applicator stick and hold it in your primary hand (right hand if right-handed).
• Holding the test tube in you other hand with palm and lower fingers, remove the cap of the test tube with your thumb and index finger. Dip the inoculated end of the applicator into the sterile medium and, if necessary, wipe the end onto the side of the tube.
• Briefly flame the end of the test tube and replace the cap.
• Always inoculate a dummy tube with no cells to verify that the medium, tubes, and sticks were sterile.
• If inoculating large cultures, dilute an existing culture 1:100.
• Grow culture for 12 to 18 hours. After 24-30 hours,

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Time required:

Cells are grown 6 hours to overnight; a total of less than 5 minutes bench time for each strain.

Procedure:

Day 1

1. Inoculate a 15 ml culture tube containing 5 ml of LBM or LBM+antibiotic selective medium with a freshly grown isolated colony. Incubate at 37 degrees C until culture is in late log or stationary phase (usually 5 hours to overnight).

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Typically, an equal volume of TE-saturated phenol is added to an aqueous DNA sample in a microcentrifuge tube. The mixture is vigorously vortexed, and then centrifuged to enact phase separation. The upper, aqueous layer carefully is removed to a new tube, avoiding the phenol interface and then is subjected to two ether extractions to remove residual phenol. An equal volume of water-saturated ether is added to the tube, the mixture is vortexed, and the tube is centrifuged to allow phase separation. The upper, ether layer is removed and discarded, including phenol droplets at the interface. After this extraction is repeated, the DNA is concentrated by ethanol precipitation.

Protocol

1. Add an equal volume of TE-saturated phenol to the DNA sample contained in a 1.5 ml microcentrifuge tube and vortex for 15-30 seconds.

• Centrifuge the sample for 5 minutes at room temperature to separate the phases.
• Remove about 90% of the upper, aqueous layer to a clean tube, carefully avoiding proteins at the aqueous:ph

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1. Warm plates to room temperature before use. Cold plates causes the top agar to solidify irregularly. DO not warm plates to 37° as the top agar will take forever to solidify.

• Prepare top agar as the appropriate liquid medium with 0.7% agar. Keeping 100 mL bottles is convenient. For phages, use λ top agar, which is less rich and yields bigger plaques.
• Melt top agar in the microwave completely. Allow the agar to boil after liquification; incompletely melted agar looks liquid, but is grainy.
• Allow agar to cool to 48°. Store in the 48° waterbath for convenience; old agar turns dark and should be discarded. Agar with drugs cannot be stored.
• Add cells and phage to a 13 mm yellow-capped tube in the 48° temperature block.
• Remove the cap and add 2.5 mL molten top agar. Do not replace cap.
• Immediately vortex the mixture, flame the top, and dispense the mixture onto the appropriate plate. Swirl the plate to evenly distribute the agar.
• Allow the plate to stand for several minutes as the agar cools.
• Aft

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Time required:

• Day 1: Overnight
• Day 2: Overnight
• Day 3: 4 hours to grow culture
• 2 hours to prepare the competent cells

Procedure:

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Author: Biotechniques

Polar lipids are generally extracted from dry cell material using chloroform:methanol:0.3% NaCl (1:2:0.8 v/v/v). This may be carried out by adding 9.5 ml of this mixture to 100 mg of freeze dried cells, or by adding a suitable amount of chloroform, methanol and 0.3% NaCl to the cell material, or to the aqueous methanolic phase remaining from the lipoquinone extraction.

1. The aqueous methanolic phase (4 ml total volume), together with the cell material from the lipoquinone analysis, is diluted with 5.5 ml of Chloroform:Methanol (2.5:3.0 v/v) to give a chloroform, methanol, 0.3% NaCl (1:2:0.8 v/v/v) mixture.

• The mixture is placed in a 15 ml bottle with a teflon lined screw cap (check that the magnetic stirrer is in the bottle), gassed briefly with nitrogen, sealed and heated for 15 min at 80 C (with occasional shaking).
• Allow the mixture to cool to room temperature on a magnetic stirrer. Check that the mixture is homogeneous, the presence of excess hexane will cause phase separ

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Authors: Greenfield Sluder, Joshua J. Nordberg, Frederick J. Miller and Edward H. Hinchcliffe

This protocol was adapted from “A Sealed Preparation for Long-Term Observations of Cultured Cells,” Chapter 18, in Live Cell Imaging (eds. Goldman and Spector). Cold Spring Harbor Laboratory Press, Cold Spring Harbor, NY, USA, 2005.

INTRODUCTION

The continuous long-term observation of cultured cells on the microscope has always been a technically demanding undertaking. This protocol describes a sealed preparation that allows the continuous long-term observation of cultured mammalian cells on upright or inverted microscopes without environmental CO2 control. The preparation allows for optical conditions consistent with high-quality imaging and good cell viability for at least 100 hours. The preparation is an aluminum support slide with a square aperture cut in its center. The coverslip bearing the cells is attached to the top of the slide with a thin layer of silicone grease, and the bottom of the slide i

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1. Sterilize velvet squares with widths at least 50% longer than the diameter of a Petrii plate.

• Streak strains to be printed for single colonies.
• Prepare a printing master plate. Place a grid sheet under the base of a sterile plate. Typically, the grids contain 24 or 50 squres, but grid between 9 and 121 squares have been used. Use a grid system of the appropriate size for the number of strains to be printed.
• Pick a single colony of each strain and hatch an 'X' shape in the middle of a grid square. Note which grid contains what strain. Make 'X' shapes rather small since they tend to get larger on subsequently printed plates. Include control strains on the last few grid-squares.
• Grow the master plate overnight.
• Prepare media for printing. Each plate should be dried so that the velvet is not excessively wet after printing.
• Place a sterile velvet face up on a print column and fix the sides with a collar. Print the master plate onto the velvet surface pressing lightly but evenly to transfer all patc

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Authors: Joachim Weischenfeldt and Bo Porse1

Corresponding author: ([email protected])

INTRODUCTION

Bone marrow-derived macrophages (BMM) are primary macrophage cells, derived from bone marrow cells in vitro in the presence of growth factors. Macrophage colony-stimulating factor (M-CSF) is a lineage-specific growth factor that is responsible for the proliferation and differentiation of committed myeloid progenitors into cells of the macrophage/monocyte lineage. Mice lacking functional M-CSF are deficient in macrophages and osteoclasts and suffer from osteopetrosis. In this protocol, bone marrow cells are grown in culture dishes in the presence of M-CSF, which is secreted by L929 cells and is used in the form of L929-conditioned medium. Under these conditions, the bone marrow monocyte/macrophage progenitors will proliferate and differentiate into a homogenous population of mature BMMs. The efficiency of the differentiation is assessed using fluorescence-activated cell sorting (FAC