Collagen extraction from pretreated bone/tooth samples

Prepare twenty 50-mL (or similar size) glass beakers, one per sample and one per each internal standard. Label them with respective sample IDs (or internal standard IDs with a date of the extraction starting day) symbols using a water-proof marker.

Weigh each beaker using the lab balance (±0.01 mg); note the weight in the Collagen extraction protocol form.

Place samples (previously cleaned, defatted and crushed to <0.7 mm powder) and internal standards (pre-treated likewise) into their respective beakers. Carefully check the ID of each sample and internal standard and each beaker to not misplace the samples nor the standards.

Weigh each beaker with a sample using the lab balance (±0.01 mg); note the weight in the Collagen extraction protocol form.

Place one magnetic stir bar inside each beaker.

Fill the “1M HCl” jug with 1M HCl (600 mL); fill the “0.125M NaOH” jug with 0.125M NaOH (600 mL).

Fill the “0.125M NaOH” wash bottle with 0.125M NaOH (usually around 200 mL is enough); fill the “water” wash bottle with Milli-Q water (usually around 200 mL is enough).

Join the membrane filter holder (with bottle attached) with the pump via the rubber pipe; make sure that the set is working.

Place each of 4 stopwatches near each of 4 magnetic stirrers; make sure that all stopwatches are ready to count down 20 minutes.

Switch on the magnetic stirrers.

Add 30 ml of 1M HCl from the “1M HCl” jug to the first beaker, place the beaker immediately on the magnetic stirrer and switch the stopwatch on for 20 minutes.

In the meantime, observe the reaction, note in the Collagen extraction protocol form the color of the residuum and the color, transparency, and any unusual characteristics (such as bubbling) of the liquid.

In the meantime, place the 5-µm nitrocellulose filter on the membrane filter holder using clean metal tweezers (must be cleaned with Milli-Q water anytime it touched anything else than the filter). Wet the filter slightly with the Milli-Q water using the "water" wash bottle. OPTIONAL: note the alternative workflow in step 20.

Switch on the pump. Make sure that the filter adheres well to the holder (it is hold well by the vacuum).

When 20 minutes counting down is over, immediately take off the beaker from the magnetic stirrer, remove the magnetic stir bar from the beaker using the magnetic rod; if any bone powder has stacked to the bar, rinse it back to the beaker with 0.125M NaOH using the “0.125M NaOH” wash bottle.

Carefully pour out the liquid and the powder from the beaker on the filter; wash down any powder which stayed in the beaker with 0.125M NaOH using the “0.125M NaOH” wash bottle; all steps 15-17 should take no longer than 30 seconds.

Wait several seconds until the pump has sucked off all the liquid; the sample powder should still seem wet, in other case it can jump out with easy; if powder seems dry, wet it by careful rinsing it with 0.125M NaOH using the “0.125M NaOH” wash bottle.

Switch off the pump.

Collect the remaining powder using the clean metal spatula and place it again in its glass beaker. It is recommended to place the filter on the beaker’s inner wall and scratch down all the powder into the beaker. Any ripped-off fragments of the filter should also go to the beaker.

Remove and dispose the filter using the tweezers or a spatula (use clean tools for each sample - replace them or clean them with Milli-Q water between each samples); you may immediately place a new 5-µm filter on the membrane filter holder using clean tweezers or clean spatula and wet it slightly with Milli-Q water using the "water" wash bottle; if so, you will skip step 13 for the next sample.

If the bottle attached to the membrane filter holder is full, dispose the liquid (follow the local regulations for disposing chemicals).

Add ml of 0.125 M NaOH from the “0.125M NaOH” jug to the beaker and tightly secure the beaker’s opening with a piece of parafilm (it is to prevent the absorption of atmospheric CO₂ by the NaOH solution). Note the time and date in a Collagen extraction protocol form.

Repeat steps 11-22 for all samples and internal standards. It is recommended to do the procedure for each sample in 5-minute intervals: start the next sample’s procedure when the previous sample’s stopwatch shows exactly 15 minutes left). In this way, you will effectively use all four stirring stations. It is possible to do it in 4-minute intervals, but this needs more experience. It is also possible to do it >5-minute intervals, which is recommended for beginners; however, the total procedure will take then more time.

Leave the beakers for 20 h.

OPTIONAL: The beakers may be placed on a shaking plate. If so, make sure that all beakers are placed safely. Switch the shaking plate on and adjust the speed. The shaking speed depends on the beaker size and amount of NaOH, and should ensure good mixing of NaOH. Usually the proper speed is between 100 rpm and 150 rpm. The shaking plate is to be switched off before the step 33.

Switch off the magnetic stirrers and the pump.

Wash the equipment (the membrane filter holder, the spatula, the tweezers, the jugs, magnetic stir bars); you may use tap water, but at the end rinse all tools with Milli-Q water several time; leave them for drying.

Clean up the working space.

Note in the Collagen extraction protocol form the color of the residuum and the color, transparency, and any unusual characteristics (such as bubbling) of the liquid for each beaker.

Mark 15-mL glass test tubes with your sample IDs and internal standard IDs.

Fill the “pH=2.00 HCl” wash bottle with pH=2.00 HCl, and fill the “water” wash bottle with Milli-Q water.

Start this step exactly 20 h after the first beaker was filled with NaOH.

Join the membrane filter holder (with bottle attached) with the pump via the robber pipe; make sure that the set is working; place the 5-µm filter on the membrane filter holder using the clean tweezers. Wet the filter slightly with the Milli-Q water using the "water" wash bottle.

Switch on the pump. Make sure that the filter adheres well to the holder (it is hold well by the vacuum).

Pour out the liquid and the bone powder from the beaker (follow the previous day’s order of samples) on the filter; use the “water” wash bottle with Milli-Q water to rinse the beaker and to wash down any powder which stacked in the beaker.

Wait several seconds until the pump has sucked off all the liquid; bone powder should still seem wet, in other case it can jump out with easy; if bone powder seems dry, wet it by careful rinsing with Milli-Q water using the “water” wash bottle. Switch the pump off.

Collect the remaining powder using the clean metal spatula and place it in a relevant 15-mL glass test tube (OPTIONAL: if some powder has stacked to the filter, place also the entire filter in the test tube); make sure that the sample ID or internal standard ID on the beaker matches that on the test tube.

Add around 5 mL of a pH=2.00 HCl to the test tube (it can be more, but not more than a half of the tube’s height) using the “pH=2.00 HCl” wash bottle (it is recommended to check the 5-mL level on a test tube using a pipette and then mark it at the same elevation on all tubes); make sure the all the powder (OPTIONAL: and the filter, if inserted) is covered by the liquid; if not, shake the tube carefully until all the powder (OPTIONAL: and the filter, if inserted) has dropped into the liquid.

Close the test tube tightly with a cup.

Place the closed test tube with the powder on a test tubes’ metal rack.

Remove and dispose the filter (if remained on the holder) using the tweezers, and place new 5 µm filter on the membrane filter holder.

If the bottle attached to the membrane filter holder is full, dispose the liquid (follow the local regulations for disposing chemical reagents).

Repeat steps 32-40 until all samples and internal standards are inside the test tubes; try to keep ~5 minutes interval (or the interval that you kept on the previous day) between starts of samples, so that each sample starts ~20 h after it has been placed on the shaking plate.

Place the rack with test tubes in an oven, switch on the oven and set the temperature to 100°C for 17 h; note the time and date in a Collagen extraction protocol form.

Wash off the symbols from the beakers using lab paper and acetone.

Clean up the working space; wash the membrane filter holder and its bottle, the beakers, the tweezers, and the jugs; you may use tap water, but at the end rinse all tools with Milli-Q water several time; leave them for drying.

Prepare twenty 10-mL LSC vials, one per sample and one per each internal standard. Label them with respective sample IDs (or internal standard IDs with a date of the extraction starting day) symbols using a water-proof marker; write each symbol twice: once at the side of the vial and separately at the cap; secure the symbol on the glass with a piece of a transparent scotch.

Weigh each LSC vial (without a cap) using the lab balance (±0.01 mg); note the weight in the Collagen extraction protocol form.

Start this step exactly 17 h after placing the tubes in the oven.

shaking plateSwitch off the oven and take the rack with the test tubes out (be careful; the oven door, the oven interior, the rack, and the test tubes are hot!).

Let the test tubes cool down to the room temperature. OPTIONAL: if needed, samples can be stored in a freezer for several days before proceeding to the next step.

Clean well the bottle of the membrane filter holder by rinsing it several times with Milli-Q water.

Join the membrane filter holder (with bottle attached) with the pump via the rubber pipe; make sure that the set is working; make sure that the bottle is clean and is attached (important! The bottle will collect the sample); place the 5-µm filter on the membrane filter holder using the tweezers.

OPTIONAL: if the size of the filter holder allows, the LSC vial may be attached to the holder instead of the holder's bottle. If so, you will collect the supernatant right away in its final LSC vial, not in the bottle. The step 56 is then modified accordingly.

Switch on the pump.

Pour out the liquid from the test tube on the filter; use the “water” wash bottle with Milli-Q water to wash down carefully any remaining liquid – pour all the liquid from the test tube onto the filter (OPTIONAL: if you placed the filter into the test tube on the previous day, rinse this old filter with Milli-Q water holding it above the membrane filter holder with tweezers).

Wait several seconds until the pump has sucked off all the liquid.

Switch off the pump.

Carefully detach the bottle from the membrane filter holder.

Collect all the liquid from the bottle into the relevant 10-mL LSC glass vial (make sure that the sample ID or internal standard ID on the test tube matches that on the LSC vial); use "water" wash bottle with Milli-Q water to wash out carefully any remaining liquid, but try not to add too much water – the entire collected liquid should not exceed a half of the 10-mL LSC vial’s volume (OPTIONAL: if there is drastically more liquid than around a half of a LSC vial, then you need to split the liquid down into two vials as follow: take a new LSC vial, mark it with the same ID adding "part b", protect the ID symbol with a piece of transparent scotch, weigh the vial (without a cap) and note its weight in the "notes" cell of the Collagen extraction protocol form; transfer an excessing portion of a liquid into this vial; notice that "the extract placed in two (or more) vials, numbers … (note both IDs)" in the "notes" cell of the Collagen extraction protocol form).

Close the LSC vial(s) tightly with a relevant cap (each time check if the ID on the vial matches that on the cap).

Clean the bottle of the membrane filter holder well by rinsing it at least 3 times with Milli-Q water (using the “water” wash bottle).

Remove all the water from the bottle (you may use compressed air if necessary; small amount of water left inside is acceptable).

Note any observed changes to the filter, such as its color, in the "notes" cell of the Collagen extraction protocol form. Remove and dispose the filter using the tweezers.

Clean the membrane holder and the funnel (if you used it) by rinsing it at least 3 times with Milli-Q water (using the “water” wash bottle).

Attach the clean bottle to the clean membrane filter holder.

Place new 5-µm filter on the membrane filter holder.

Repeat steps 51-63 until all samples and internal standards are inside their 10-mL LSC vials.

Place the LSC vials in a plastic container; check again if all vials are tightly closed.

Place the plastic container with the vials in a freezer, set the temperature at -30°C (or lower, if possible); make sure that the freezer is working; note the time and date in a Collagen extraction protocol form. Leave the vials in the freezer for at least 2 days.

Clean up the working space; wash the membrane filter holder and its bottle, the tweezers and the jugs; you may use tap water, but at the end rinse all tools with Milli-Q water several time; leave them for drying.

Take the container with the vials out from the freezer. Make sure that all samples are frozen (only ice is visible, no liquid water) – if not, place them again in the freezer for another 1 day.

Take off all the caps; place the caps on a clean plastic tray and keep in a safe and clean place (it is recommended to cover the tray with a large piece of aluminum foil, or to use a plastic box with a lid instead of a tray).

Cover tightly the opening of each LSC vial with around 2 cm by 2 cm piece of aluminum foil (keep the foil pieces on another clean plastic tray).

Use a clean metal lancet or awl to make several holes in the aluminum foil (to allow the air and vapor escaping from the vial during drying).

Place the vials again in the container and place the container with the vials again in the freezer; try to execute all steps 68-72 quickly, to not allow the liquid melting down.

Leave the vials in the freezer at -30°C for at least 2 h, to ensure that the liquids are completely frozen and cold.

Switch the freeze-dryer on and set up the 48-h collagen drying program, following the freeze-dryer’s instruction (minimum requirements for the program include: -55°C or lower condenser temperature; 0.01 mBar or lower pressure; these conditions kept for at least 48 h; heating the shelves at +20°C, starting not earlier than the -55°C and 0.01 mBar conditions are obtained).

When freeze-dryer is ready to work and its program is in “loading” phase, take the container with the vials off from the freezer.

Immediately place vials on the freeze-dryer’s shelf (shelves).

OPTIONAL: Repeat steps 75-76 for any other series of samples ready for freeze-drying.

Place a freeze-dryer’s cylinder over freeze-dryer’s chamber and shelves; make sure that the cylinder adheres well to the rubber gasket; lock all external valves of the freeze-dryer (such as an air-valve and a water-drain-valve).

Switch the freeze-dryer to the next phase of drying, following the freeze-dryer’s instruction; try to execute all steps 75-79 quickly, to not allow the ice melting down. Make sure that the freeze-dryer follows the program; follow the freeze-dryer’s instruction in the case of problems.

When freeze-drying is over, follow the freeze-dryer’s instruction to switch the freeze-dryer off, to open the cylinder and to drain the water off.

Make sure that the samples and internal standards look fluffy and cotton-like (if not, you need to add few mL of Milli-Q water to a vial and shake it carefully for few minutes to allow all the collagen dissolve, freeze the sample again and repeat the entire freezing and freeze-drying procedure).

Immediately take the vials off of the freeze-dryer.

Immediately remove the aluminum foil covers and dispose them; be careful to not lose any collagen.

Quickly close the vials tightly with their relevant caps (each time check if the ID on the vial matches that on the cap); be careful to not lose any collagen. Use a mouth-covering mask (such as an anti-dust mask) to prevent breathing into the vials, to not contaminate nor blow out the collagen.

Immediately place the vials in the plastic container.

OPTIONAL: Repeat steps 83-86 for any series of samples which was freeze-dried.

Weigh each vial (without a cap) with the collagen inside, using the lab balance (±0.01 mg); note the weight in the Collagen extraction protocol form. Do the weighing fast to minimize the time of collagen exposure to the atmosphere, to avoid contamination and absorbing the water vapor by the collagen. It is recommended to do this step during a sunny, low-humidity day.

Immediately after the weighing close the vial tightly with its cap and place it in its plastic container.

Place the container with the samples in the desiccator cabinet. Upon dry and cool conditions (room temperature or below, humidity <5%) the sample can be stored for years.

Calculate the collagen yield using the Collagen extraction protocol form (yield [in %] = [weight of LSC vial with freeze-dried collagen – weight of empty LSC vial] / [weight of beaker with bone powder – weight of empty beaker] * 100%). If the attached Collagen extraction protocol form file is used, it will calculate the yield automatically.