An Improved Method for Collection of Cerebrospinal Fluid from Anesthetized Mice

Podsumowanie

This protocol describes an improved technique for the abundant collection of cerebrospinal fluid (CSF) with no contamination from blood. With greater sample collection and purity, more analyses can be performed using CSF to further our understanding of diseases that affect the brain and spinal cord.

Streszczenie

The cerebrospinal fluid (CSF) is a valuable body fluid for analysis in neuroscience research. It is one of the fluids in closest contact with the central nervous system and thus, can be used to analyze the diseased state of the brain or spinal cord without directly accessing these tissues. However, in mice it is difficult to obtain from the cisterna magna due to its closeness to blood vessels, which often contaminate samples. The area for CSF collection in mice is also difficult to dissect to and often only small samples are obtained (maximum of 5-7 µL or less). This protocol describes in detail a technique that improves on current methods of collection to minimize contamination from blood and allow for the abundant collection of CSF (on average 10-15 µL can be collected). This technique can be used with other dissection methods for tissue collection from mice, as it does not impact any tissues during CSF extraction. Thus, the brain and spinal cord are not affected with this technique and remain intact. With greater CSF sample collection and purity, more analyses can be used with this fluid to further aid neuroscience research and better understand diseases affecting the brain and spinal cord.

Wprowadzenie

The CSF is a valuable body fluid for analysis in neuroscience research. The CSF is primarily made from blood plasma, containing few cells (no red blood cells and only a few white blood cells) and is almost protein-free. It is one of the fluids in close contact with the central nervous system (CNS) and it can pass many electrolytes out from the brain and spinal cord to the peripheral system. In humans, CSF samples can be collected to aid in diagnosing disease or for research purposes in clinical trials, as a spinal tap (or lumbar puncture) is a minor, invasive procedure: the CSF fluid can reflect changes in the CNS without having to directly access these tissues. Thus, in recent years, for research purposes in the clinic, CSF samples have been obtained from patients of neurodegenerative diseases such as Alzheimer's disease and other dementias^1,2,3. There are many biomarker assays which have been developed using CSF samples to potentially aid in diagnosing diseases in the clinic^2,3. However, there is much debate on the reliability of these assays to produce consistent, sensitive results to specifically diagnose disease^4,5. So, there is a great need for the development of better assays and targets, which can be found in the CSF, to aid in producing a standard technique to diagnose neurodegenerative diseases with greater sensitivity and specificity. Due to the potential importance of human CSF samples in disease, the collection of CSF from rodents in neuroscience research is also of interest.

Mice are important animals in biological and medical research and allow for the testing of potential therapeutic compounds and proof-of-concept studies before human clinical trials. However, in mice it is difficult to obtain CSF samples due to its closeness to the brain in a small animal, as the usual method of CSF collection in mice is to obtain it via the cisterna magna, an opening between the cerebellum and dorsal surface of the medulla oblongata. This causes difficulty in collecting CSF samples as this area is difficult to dissect to and in close proximity to blood vessels, increasing the risk of contamination from blood cells. Due to these difficulties, most researchers can only obtain a small amount of CSF for analysis (usually stated as 5-7 µL) and the contamination of CSF samples by blood cells is a primary concern for analyses^6,7,8,9. Blood contamination can obscure results and not truly reflect the state of the CNS. Furthermore, limited sample collected can impact research as the usual amount collected from mice is enough for only one measurement (in duplicate or triplicate) using enzyme-linked immunosorbent assay (ELISA). Thus, CSF samples are usually pooled from multiple mice in order to have enough sample to run multiple assays. Developing a protocol for the abundant, uncontaminated collection of CSF from mice is greatly desired and will be beneficial in improving neuroscience research using rodents.

In this protocol, a technique for the abundant (an average of 10-15 µL) collection of CSF from anesthetized mice is described in detail and improves on a currently known method of CSF collection to minimize contamination from blood¹⁰. A robust protocol for CSF collection will aid in the development of CSF-based biomarker assays, which could be used to aid in diagnosing disease, as well as improve research into the mechanisms that underlie diseases affecting the CNS.

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Protokół

All animal experiments were performed in accordance with the policies of the Society for Neuroscience (USA) and Fudan University (Shanghai, China) ethical committees. This procedure is for a non-survival surgery.

1. Setup of CSF Collection Apparatus

1. Pull the glass capillary (inner diameter 0.75 mm, outer diameter 1.0 mm) using a micropipette puller (as shown in Liu et al.¹⁰; sharpened capillary is shown in Figure 1).
2. Place a sharpened glass capillary into the capillary holder firmly mounted on a micromanipulator (Figure 1A).
3. Break the tip of the sharpened capillary with straight forceps under a dissecting microscope, so that the inner diameter of the broken tip is about 10-20 µm (Figure 1C).
4. Attach a thin tube and a syringe to the other end of the capillary holder, and connect the thin tube and the syringe with a three-way valve.
5. Shift the three-way valve to open and plunge the syringe to blow air from the syringe through the tubing to the glass capillary to expel any contaminants and create positive pressure in the capillary tube.
6. Repeat this a few times by dis-connecting and re-connecting the syringe to the three-way valve, and drawing the syringe up ~ 100-200 µL before the last re-connection of the syringe with the three-way valve (Figure 1B).
7. Move the micromanipulator with the glass capillary to the side to prevent damage during mouse dissection.

Figure 1. Microscope, micromanipulator, and capillary setup. (A) Microscope and micromanipulator with capillary attached setup, as well as a closer image of (B) the connected syringe and three-way valve to open (shown here) or close the piping, and (C) an unbroken (right) and broken (left) capillary tip ready for CSF collection. Once broken, the tip of the capillary should have an inner diameter of 0.75 mm, outer diameter of 1.0 mm. Please click here to view a larger version of this figure.

2. Mouse Dissection

1. Anesthetize the mouse.
   NOTE: For this protocol, C57BL/6 and amyloid precursor protein/presenilin 1 (APP/PS1) transgenic (Alzheimer's disease mouse model) mice were used and anesthetized with 4% chloral hydrate (in dH₂0) at 10 µL/g of mouse weight via intraperitoneal injection.
2. When fully anesthetized, ensure that the mouse is adequately anesthetized. The withdrawal response to toe pinch, by slightly extending the hind limbs and pinching the toes and watching for a withdrawal response, is performed to assure adequate anesthesia. Check by pinching all four limbs; if there is no reaction the mouse is properly anesthetized. Cut and trim the hair at the back of the head of the mouse from above the eyes, between the ears to the bottom of the neck using dissecting scissors (Figure 2A). Alternatively, clippers or depilatory cream can be used to aid in hair removal.
3. Secure the head of the mouse using a mouse adapter (back of the head facing up with the nose pointed down at ~ 45 °, Figure 2A). Fix ear bars between the ears and eyes of the mouse and tighten. Make sure the head cannot move and is secured tightly in the adapter by gently pushing down onto the head of the mouse.
4. Recheck withdrawal response to toe pinch prior to making surgical incision and regularly thereafter to assure surgical plane of anesthesia. Visually note any change in respiratory rate during the procedure to indicate changes in planes of anesthesia. A clean, non-aseptic, technique can be used for this short non-survival surgery. Using scissors and curved forceps, cut through the skin and the first layer of muscle until the base of the skull is exposed with only a thin layer of muscle.
   NOTE: Cut through the mouse skin across the back of the neck and then cut the skin down the middle of the skull between the ears and eyes of the mouse. Skin and tissue can then be pulled apart and out of the area over the cisterna magna.
5. While viewing through the dissecting microscope, carefully dissect away the last thin layers of muscle over the base of the skull using forceps and move these layers of muscle to the side and out of the area of interest. If there is bleeding, use cotton buds to remove and help stop the bleeding.
6. Once the dura over the cisterna magna is exposed (triangular in shape with usually 1-2 large blood vessels running through the area; either side of or between the blood vessels is optimal for capillary insertion and CSF collection, Figure 2B), use a wet cotton bud to wipe the membrane clean and then use a dry cotton bud to dry the area.

Figure 2. Mouse and dissection setup with representative images of capillary insertion for CSF collection. Setup of mouse ready for CSF extraction with (A) shaved head clamped in place for dissection and (B) detailed image (10x view) of a dissected mouse dura covering the cisterna magna (dashed arrow shows a blood vessel running through the area and solid arrow shows area optimal for capillary insertion). Detailed images (10x view) of (C) sharpened tip of glass capillary aligned against dura of cisterna magna, (D) the point at which the capillary almost punctures the dura, with some resistance from dura, and (E) the capillary tip tapped through the dura to collect CSF. CSF should be a clear liquid collected in the glass capillary (dotted arrow). All scale bars at the bottom right of each microscope image indicate 100 µm. Please click here to view a larger version of this figure.

3. CSF Collection from Anesthetized Mouse

1. Align the glass capillary to the back of the mouse head so that the sharpened point is just behind the membrane at a ~ 30-45 ° angle (Figure 2C).
2. Plunge the syringe once or a few times and draw the syringe up ~ 100-200 µL (to make sure there are no contaminants and there is negative pressure in the glass capillary).
3. Using the micromanipulator, move the capillary tip closer to the membrane until resistance can be seen, but not puncturing it (Figure 2D).
4. Once resistance is seen between the tip of the capillary and the membrane, gently tap the capillary tube through the membrane by knocking on the micromanipulator controls. Use the microscope to see the sharpened point of the capillary puncture into the cisterna magna. CSF should automatically be drawn into the capillary tube once the opening has been punctured (Figure 2E).
5. Leave the capillary to slowly collect CSF. If the CSF has stopped drawing out, draw the syringe up a small amount slowly (~ 50 µL) to create more negative pressure in the capillary to extract out the CSF.
   NOTE: Alternatively, the capillary can be gently and slowly pulled out of the brain using the fine micromanipulator controls to allow CSF to flow into the capillary again. A total amount of ~ 10-15 µL of CSF (~ 3-4 cm in the capillary) takes ~ 10-30 min, and occasionally 20 µL or more can be collected. Although not absolutely necessary, it is recommended to use a heat pad during CSF collection. CSF collection was performed at room temperature (~ 23 °C).
6. Once the amount of desired CSF is collected, close the tubing by shifting the three-way valve to close (to stop drawing out the CSF).
7. Take out the syringe connected to the tubing, and open and then close the tubing (to create equalized pressure in the capillary, tubing, and the area outside the capillary). Reconnect the syringe with the three-way valve, but do not plunge the syringe.
8. Gently remove the glass capillary from the mouse using the micromanipulator.
9. Place the collection tube (1.5 mL microtubes with 1 µL of 20x protease inhibitor) under the sharpened point of the glass capillary.
10. Open the tubing and gently plunge the syringe: the CSF should flow out of the capillary and into the collection tube.
11. After collection, quickly centrifuge (pulse spin for 5 s at maximal speed using a mini centrifuge) the CSF to mix the sample with the protease inhibitor at the bottom of the collection tube.
    NOTE: CSF samples can be aliquoted and then stored for further analysis at -80 °C. The rest of the mouse can be dissected for other tissues, such as brain and spinal cord. The mouse is still alive at this stage and thus blood can also be collected, if needed.

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Wyniki

Using the procedure outlined here (Figure 1 and Figure 2), the CSF immediately collected in the capillary should be clear (Figure 2E), not pink or red. If there is a pink to red tinge to the fluid collected in the capillary, then there was contamination with blood.

As an example of the application of the CSF sample collected with this metho...

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Dyskusje

This protocol describes in detail a technique which improves on current methods¹⁰ of CSF collection to minimize contamination from blood and allow for the abundant collection of CSF (on average ~ 10-15 µL can be obtained) from mice. When breaking the capillary tip, the tip of the capillary should not be too small (as then the CSF will be extracted very slowly) or too big (will not be fine enough to collect the CSF and tissue can become lodged in the capillary). Care should be taken when ...

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Materiały

Name	Company	Catalog Number	Comments
Chloral hydrate (used as anesthetic)	Sinopharm Chemicals Reagen Co. Ltd.	30037517	CAS number 302-17-0.
Dissecting scissors	66 vision technology	54002
Dissecting curved forceps	66 vision technology	53072
Dissecting straight forceps	66 vision technology	53070
Mouse adapter (with ear bars)	Made in-house.	N/A	Similar equipment available from World Precision Instruments.
Dissecting microscope	Meiji Labax	Model 15381
Micromanipulator	World Precision Instruments	M3301
Magnetic base for micromanipulator	Kanetec	MB-K
Glass capillaries	World Precision Instruments	1B100-4
Micropipette puller	Sutter Instruments	Model P-1000
Syringes (1ml)	Tansoole	02024692	For 1ml.
Microtubes (1.5ml)	Axygen	MCT-150-C
Protease inhibitor Cocktail Set III EDTA-free	Calbiochem	539134
Human Aβ42 ELISA kit	Invitrogen	KHB3441
Piping (teflon tubing)	World Precision Instruments	MMP-KIT	Obtained from a microinjection kit and attached to the capillary holder and syringe.
Mini centrifuge	Tiangen Biotech	OSE-MC8
Cotton buds	Obtained from any household store/pharmacy.	N/A