National Institutes of Health grant DK124619 to KHC.
Start-up funds and pilot research award, Department of Medicine, University of Rochester, NY, to KHC.
The Del Monte Institute for Neuroscience Pilot Research Award, University of Rochester, to KHC.
University Research Award, Office of the Vice President for Research, University of Rochester, NY, to KHC.
MUR designed and performed the method, analyzed results, prepared graphs and figures, and wrote and edited the manuscript. KHC conceived and supervised the study, analyzed results, and wrote and edited the manuscript. KHC is the guarantor of this work. All authors approved the final version of the manuscript.

All mouse procedures were approved by the Institutional Animal Care and Use Committee at the University of Rochester and were performed according to the US Public Health Service guidelines for the humane care and use of experimental animals. Six weeks old C57BL/6J male mice used for this study were commercially obtained. All the animals were group housed (5 mice per cage) in a room with a 12 h day/night cycle and were given access to food and water ad libitum. After the mice were implanted with a cannula for infusing glucose into the lateral ventricle, they were single housed to prevent any damage to the implants from other mice.
1. Assembly of osmotic minipumps
<ol>
	<li>Prepare artificial cerebrospinal fluid (aCSF) according to the following protocol. Dissolve NaCl (8.66 g), KCl (0.224 g), CaCl2.2H2O (0.206 g), and MgCl2.6H2O (0.163 g) in 500 mL of sterile water to prepare solution A. Dissolve Na2HPO4.7H2O (0.214 g) and NaH2PO4.H2O (0.027 g) in 500 mL of sterile water to prepare solution B. Then, mix solution A and B in a 1:1 ratio to prepare the aCSF. 
	NOTE: This formulation of aCSF contains no glucose.</li>
	<li>To prepare the 50% glucose solution, add 50 g of glucose to 50 mL of aCSF in a beaker. Place the beaker on a hot plate and bring the temperature of the suspension to 60&#160;&#176;C. Mix the suspension with a magnetic stirrer until the glucose is completely dissolved.</li>
	<li>Add aCSF to the beaker to make the final volume 100 mL. Pass the solution through 0.22 &#181;m pore filter to sterilize it.</li>
	<li>Prepare a 0.9% saline (NaCl) solution and pass the solution through the 0.22 &#181;m filter.</li>
	<li>Place all the components of the minipump and infusion kit in a sterile tray and cut the tubing from the brain infusion kit to 1 inch in length with a sterile scissor. Hold the tubing with a sterile hemostat and push it on the top of one end of the cannula. Connect the other end of the tubing to the top side of the flow modulator. Cover both ends of the tubing with a small amount of glue for a stronger connection.</li>
	<li>Attach a 27 G needle to a 1 mL syringe and fill it with either aCSF or the glucose solution. Insert the 27 G needle into a small piece of tubing. Then, connect the tubing to the open end of the flow modulator and fill it with the corresponding solution until drops of the solution start coming out from the cannula. Remove any air bubbles trapped in the tubing.</li>
	<li>To fill the minipump, hold the pump upright and insert the syringe needle into the pump. Fill it with the corresponding solution until a drop of the solution starts forming at the opening of the minipump.</li>
	<li>Slowly insert the flow modulator into the pump while making sure not to introduce any air bubbles in the assembly.</li>
	<li>After the osmotic minipumps are assembled, transfer them to a 50 mL conical tube. Fill the tube with the sterile saline so that the pumps are completely submerged, while making sure that the cannula stays out of the saline solution. Four such assembled pumps can be placed in one 50 mL tube.</li>
	<li>Loosely place the cap on each tube and incubate the pumps at 37&#160;&#176;C to prime for at least 48 h.</li>
</ol>
2. Surgery to implant osmotic pumps
<ol>
	<li>Pre-surgery procedure
	<ol>
		<li>Sterilize the surgical instruments in an autoclave and let them cool down for at least 30 min before use. The autoclave conditions involve steam sterilization at 121 &#176;C for 30 min.</li>
		<li>Disinfect the surgery area with 70% ethanol. Wipe the knobs, ear bars, and stereotaxic frame surface. 
		NOTE: Surgery should be performed in an aseptic environment, keeping in mind to avoid touching non-sterile surfaces while performing surgery. Replace gloves if there is a suspicion of contamination.</li>
		<li>Place the mouse in an anesthesia chamber to anesthetize it with 3% isoflurane mixed in air for 3-5 min. Then, remove the mouse from the chamber and place it on a sterile drape. Maintain anesthesia via a nose cone at 1.5%-2%.</li>
		<li>Pinch the mouse hind paw to observe any reflex and verify the depth of anesthesia.</li>
		<li>Inject 5 mg/kg carprofen subcutaneously for analgesia.</li>
		<li>Use clean clippers to clip the surgery area on the mouse head. Rub the scalp first with povidone-iodine solution and then with alcohol swabs thoroughly for at least 2 min. Repeat this process at least three times.</li>
		<li>Apply an eye lubricant to prevent the eyes from drying during surgery.</li>
	</ol>
	</li>
	<li>Surgery
	<ol>
		<li>Place the mouse on the stereotaxic frame and fit the head on the incisor bar. Cover the nose with the nose cone. Apply sterile drapes to secure the surgical area.</li>
		<li>Shift the flow of isoflurane to the nose cone. Continue to maintain the anesthesia at&#160;1.5%-2% isoflurane.</li>
		<li>Place a heating pad under the mouse to maintain the body temperature.</li>
		<li>Put on new sterile gloves and, with a scalpel, make a midline incision (~10-15 mm long) on the scalp. Expose the skull surface with a spatula while using cotton tips to wipe any bleeding. 
		NOTE: Use a cauterizer if bleeding profusely, which is rare.</li>
		<li>Rub with a cotton tip dipped in 3% hydrogen peroxide to expose the cranial sutures.</li>
		<li>Take note of bregma and lambda to navigate the skull coordinates. Set all the coordinates to zero at bregma. 
		NOTE: These coordinates were obtained from a mouse brain atlas14. The reference bregma-lambda distance in the atlas is 4.21 mm. To correct for age- and sex-based differences in skull size, the bregma-lambda distance for each mouse was measured and then divided by 4.21 mm to obtain a multiplication factor. For example, if the distance between bregma and lambda was 4.65 mm, division by 4.21 mm gave a multiplication factor of 1.10. All the coordinates obtained from the atlas were then multiplied with this factor to measure the actual coordinates that were used to drill the hole. In the above example, the resulting anterior-posterior (AP), and medial-lateral (ML) coordinates would be 0.55 and 1.1 mm, respectively (i.e., 1.1 x 0.5 or 1 mm).</li>
		<li>Use an electric drill to make a hole at these coordinates: 0.5 mm posterior to bregma and 1 mm lateral to midline toward the right.&#160; 
		NOTE: While drilling the hole in the skull, be careful not to rupture the dura mater. Make sure the drill bit is sterile. It can be placed with other instruments in the same autoclave pouch for sterilization.</li>
		<li>Insert a hemostat under the incised skin and push it to the place where the osmotic minipump will be implanted. Gently open and close the hemostat to make a pocket for the minipump.</li>
		<li>Use a sterile hemostat to pick&#160;a primed osmotic minipump from the tip of the flow modulator. Push the pump through the incised skin into the pocket. 
		NOTE: Be careful to not let the pump and the cannula touch any non-sterile surface.</li>
		<li>Apply some glue on the underside of the cannula and fix it in the cannula holder. Lower the cannula slowly through the drilled hole 2 mm ventral to the skull surface. Leave it there for at least 5 min to allow the glue to solidify.</li>
		<li>Before inserting the cannula down into the brain, make sure that the cannula is not clogged and that the solution is flowing properly. 
		NOTE: Sometimes, glucose may deposit at the tip of the cannula due to a very low flow rate, blocking the flow.</li>
		<li>With the help of a scissor, clip the top of the cannula making sure that it doesn&#39;t dislodge from the skull surface.</li>
		<li>Cover the cannula with a layer of dental cement for extra support.</li>
		<li>Use nylon sutures (5-0 size and 30 inches in length) to close the wound and apply topical antibiotic on the wound to prevent infection.</li>
		<li>Turn off the isoflurane and let the animal recover from the anesthesia. Then, transfer the animal to a new clean cage.</li>
	</ol>
	</li>
	<li>Post-surgery care
	<ol>
		<li>After surgery, monitor the mouse for at least 1 week.</li>
		<li>Administer 5 mg/kg carprofen subcutaneously, once every 24 hours for three days post-surgery.</li>
		<li>Let the wound heal completely before removing the sutures 7-10 days after surgery. 
		&#8203;NOTE: It was observed that mice with high CSF glucose took longer to heal. Trimming nails on mouse&#39;s hind paws minimizes the chance of injury to the site of the incision due to the scratching behavior of mice.</li>
	</ol>
	</li>
</ol>
3. Replacement of the minipumps
NOTE: Since, the minipumps used in this study last only for 4 weeks, the replacement of minipumps was also tested to extend the duration of glucose infusion, as it may be required in the case of long-term studies. This involved the following steps.
<ol>
	<li>Prepare the mice for surgery as detailed above.</li>
	<li>Make a small 1 cm vertical incision in the skin slightly above the pump.</li>
	<li>Insert a hemostat under the incised skin and pull the pump out.</li>
	<li>Remove the pump from the tubing and connect a new primed pump to the tubing.</li>
	<li>Insert the pump back inside and stitch the skin.</li>
	<li>Follow the same post-surgery care instructions as described above.</li>
</ol>
4. CSF collection procedure
<ol>
	<li>Preparation
	<ol>
		<li>Pull 1 mm diameter glass capillaries with a micropipette puller machine to obtain 0.5 mm diameter tips. The settings are: heat = 800, pull = 15, velocity = 5, and time = 200 units. Place the pulled capillaries in a sterile box till use.</li>
		<li>Attach a 27 G needle to a 1 mL syringe. Insert the pulled capillary on the needle and use a small adhesive tape to secure the connection.</li>
		<li>Firmly fix the syringe in a capillary holder of a micromanipulator.</li>
		<li>Next, prepare the mouse for surgery, as described in the pre-surgery procedure above.</li>
	</ol>
	</li>
	<li>Surgical procedure
	<ol>
		<li>Place the mouse on the stereotaxic frame. After fixing the head on the frame, as described previously, rotate the knobs to tilt the head so that the nose is facing downward. Apply sterile&#160;drapes to secure the surgical area.</li>
		<li>Make a small incision on the dorsal surface, starting from the intra-aural plane to the caudal side. 
		NOTE: Be careful not to damage the tubing while making the incision.</li>
		<li>Remove the support from the mouse body and let the mouse rest vertically so that the neck is fully extended dorsally.</li>
		<li>With the help of curved blunt forceps, gently bisect the posterior neck muscles from the midline to create a small window.&#160;Then, use a wet cotton tip applicator to gently displace the neck muscles from&#160;mid-line to&#160;the periphery.</li>
		<li>Observe whether the cisterna magna is exposed, which appears as a triangular window with a transparent dura membrane.</li>
		<li>Place the micromanipulator assembly on the stereotaxic frame next to the mouse while making sure that the capillary tip does not touch any surface.</li>
		<li>Gently break the capillary tip without disturbing the setup.</li>
		<li>While looking into the magnifying glass (magnification of 2.25x), rotate the corresponding knobs on the micromanipulator to slowly align and move the capillary tip toward the cisterna magna.</li>
		<li>Some resistance is felt once the capillary tip touches the cisterna magna membrane. Very slowly push the tip against the membrane with the help of the knobs. 
		NOTE: Be careful not to damage any blood vessels in the membrane. This step is very important to avoid any blood contamination in the CSF.</li>
		<li>Very gently pierce the membrane. CSF will start flowing into the capillary at once due to negative pressure in the capillary.</li>
		<li>Leave the setup for few minutes until ~10 &#181;L of CSF is collected in the capillary.</li>
		<li>Then, slowly draw the capillary out of the cisterna magna and gently press a sterile cotton tip applicator against the opening of the cisterna magna to stop CSF leakage.</li>
		<li>Carefully remove the capillary from the syringe and attach it to a microcap bulb dispenser. Press the bulb to transfer the CSF into a sterile microcentrifuge tube.</li>
		<li>Place a support under the mouse and rotate the stereotaxic knobs to level the head.</li>
		<li>Close the wound with nylon sutures.</li>
		<li>Inject 300 &#181;L of sterile saline subcutaneously before removing the mouse from the apparatus.</li>
		<li>Give post-operative care to the animals&#160;as described above, until it is time to remove the sutures-~7-10 days after surgery.</li>
	</ol>
	</li>
</ol>
5. Glucose assay
<ol>
	<li>Follow the protocol as described in the glucose colorimetric assay kit.</li>
	<li>Dilute the sodium phosphate buffer stock solution provided in the kit to 50 mM concentration in ultra-pure water.</li>
	<li>Prepare glucose standards in 0, 2.5, 5, 7.5, 10, 15, 20, 25, and 50 mg/dL ranges from the 1,000 mg/dL stock solution provided in the kit in diluted buffer solution.</li>
	<li>Prepare a 7 fold&#160;dilution of CSF samples in the buffer.&#160;For example, if the total CSF volume collected is 10 &#181;L, then the seven fold diluted volume will be 70 &#181;L.</li>
	<li>Pipette 15 &#181;L of glucose standards and diluted CSF in duplicates in a 96-well plate.</li>
	<li>Pipette 85 &#181;L of diluted buffer in each of the wells with standards and CSF.</li>
	<li>Add 6 mL of diluted buffer to the glucose calorimetric enzyme mixture vial and vortex it for a few seconds to mix thoroughly.</li>
	<li>Add 100 &#181;L of the enzyme mixture preparation to each well to start the reaction.</li>
	<li>Seal the plate with a cover sheet and tap on the plate gently to mix the reagents.</li>
	<li>Place the plate in an incubator at 37 &#176;C for 10 min. Vials with high glucose start to turn violet immediately.</li>
	<li>After 10 min, remove the cover and measure the absorbance at 500-520 nm using a plate reader.</li>
	<li>Calculate the glucose concentration in the CSF samples by interpolating their values from the standard curve.</li>
</ol>
6. Blood glucose assay
<ol>
	<li>Make a small lateral nick to the mouse tail using a razor blade and use a drop of blood to measure the blood glucose with a glucometer, according to the manual instructions.</li>
</ol>

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The authors declare that they have no conflict of interests.

This article reports a detailed protocol to increase CSF glucose in mice by using osmotic minipumps connected to a cannula implanted in the lateral ventricle. The chronic infusion of glucose in the mouse brain through this procedure&#160;will be useful in delineating the effects of long-term hyperglycorrhachia on cognition, systemic glucose metabolism, and energy balance&#160;and for better understanding the pathogenesis of diabetes complications.
Chronic diabetes causes brain damage that inte...

Both type 1 and type 2 diabetes impair brain function1,2,3. For example, diabetes increases the risk of cognitive decline and neurodegenerative disorders, including Alzheimer&#39;s disease3,4. Moreover, people with diabetes have defective glucose sensing in the brain5,6. This defect contributes to the pathogenesis of hypoglycemia associated unawareness and an insufficient counter-regulatory response to hypoglycemia7,8, which can be fatal if not treated immediately.
Considering that diabetes increases glucose levels in the blood as well as in cerebrospinal fluid (CSF)9, it is important to determine whether one or both of these factors contribute to impaired brain function. Whether diabetes causes brain damage by high CSF glucose alone or in combination with other factors like insulin deficiency or insulin resistance is also an open question. Animal models of type 1 and type 2 diabetes show cognitive decline and neurodegeneration in addition to an affected energy balance and peripheral glucose metabolism10,11,12,13. However, from these models, it is not feasible to uncouple the selective effects of high CSF glucose versus blood glucose levels in mediating the complications of diabetes on brain function.
This protocol describes methods to develop a mouse model of hyperglycorrhachia to test the effects of chronically high CSF glucose levels on brain function, energy balance, and glucose homeostasis. The mouse model developed through this technique presents a tool for studies investigating the etiological role of dysregulated glucose homeostasis on neural and behavioral function.
Therefore, the proposed approach will be useful in understanding the direct effects of elevated CSF glucose levels in various pathophysiological conditions.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>0.22 &#181;m syringe filter</td>
			<td>Membrane solutions</td>
			<td>SFPES030022S</td>
			<td></td>
		</tr>
		<tr>
			<td>1 mL sterile Syringe (Luer-lok tip)</td>
			<td>BD</td>
			<td>309628</td>
			<td></td>
		</tr>
		<tr>
			<td>1 mL TB syringe</td>
			<td>BD</td>
			<td>309659</td>
			<td></td>
		</tr>
		<tr>
			<td>100 mL Glass beaker</td>
			<td>Fisher&#160;</td>
			<td>N/a</td>
			<td></td>
		</tr>
		<tr>
			<td>100% Ethanol (Koptec)</td>
			<td>DLI</td>
			<td>UN170</td>
			<td>Use 70% dilution to clean the surgery area</td>
		</tr>
		<tr>
			<td>50 mL conical tube</td>
			<td>Fisher&#160;</td>
			<td>N/A</td>
			<td></td>
		</tr>
		<tr>
			<td>Allignment indicator</td>
			<td>KOPF</td>
			<td>1905</td>
			<td></td>
		</tr>
		<tr>
			<td>Alzet brain infusion kit</td>
			<td>DURECT</td>
			<td>Kit # 3; 0008851</td>
			<td>Cut tubing in the kit to 1 inch length</td>
		</tr>
		<tr>
			<td>Alzet osmotic pump</td>
			<td>DURECT</td>
			<td>2004</td>
			<td>Flow rate 0.25 &#181;L/h</td>
		</tr>
		<tr>
			<td>Anesthesia system</td>
			<td>Kent Scientific</td>
			<td>SomnoSuite</td>
			<td></td>
		</tr>
		<tr>
			<td>Betadine solution</td>
			<td>Avrio Health</td>
			<td>N/A</td>
			<td></td>
		</tr>
		<tr>
			<td>CaCl2 . 2H2O</td>
			<td>Fisher&#160;</td>
			<td>C79-500</td>
			<td></td>
		</tr>
		<tr>
			<td>Cannula holder</td>
			<td>KOPF</td>
			<td>1966</td>
			<td></td>
		</tr>
		<tr>
			<td>Centering scope</td>
			<td>KOPF</td>
			<td>1915</td>
			<td></td>
		</tr>
		<tr>
			<td>Dental Cement Liquid</td>
			<td>Lang Dental</td>
			<td>REF1404</td>
			<td></td>
		</tr>
		<tr>
			<td>Dental cement Powder</td>
			<td>Lang Dental</td>
			<td>REF1220-C</td>
			<td></td>
		</tr>
		<tr>
			<td>D-glucose&#160;&#160;</td>
			<td>Sigma</td>
			<td>G8270</td>
			<td></td>
		</tr>
		<tr>
			<td>Electric drill</td>
			<td>KOPF</td>
			<td>1911</td>
			<td>While drilling a hole avoid rupturing dura mater</td>
		</tr>
		<tr>
			<td>Eye lubricant (Optixcare)</td>
			<td>CLC Medica</td>
			<td>N/A</td>
			<td></td>
		</tr>
		<tr>
			<td>Glass Bead sterilizer (Germinator 500)</td>
			<td>VWR</td>
			<td>101326-488</td>
			<td>Place instruments in sterile water to let them cool before surgery</td>
		</tr>
		<tr>
			<td>Glucose Assay Kit</td>
			<td>Cayman chemical</td>
			<td>10009582</td>
			<td></td>
		</tr>
		<tr>
			<td>H2O2</td>
			<td>Sigma</td>
			<td>H1009-500ml</td>
			<td>Apply 3% H2O2 on skull surface to make the cranial sutures visible.</td>
		</tr>
		<tr>
			<td>Hair Clipper</td>
			<td>WAHL</td>
			<td>N/A</td>
			<td></td>
		</tr>
		<tr>
			<td>heating pad</td>
			<td>Heatpax</td>
			<td>19520483</td>
			<td></td>
		</tr>
		<tr>
			<td>Hemostat</td>
			<td>N/A</td>
			<td>N/A</td>
			<td></td>
		</tr>
		<tr>
			<td>Isoflurane (Fluriso)</td>
			<td>Zoetis</td>
			<td>NDC1385-046-60</td>
			<td></td>
		</tr>
		<tr>
			<td>KCl</td>
			<td>VWR</td>
			<td>0395-500g</td>
			<td></td>
		</tr>
		<tr>
			<td>Magnetic stand</td>
			<td>WPI</td>
			<td>M1</td>
			<td></td>
		</tr>
		<tr>
			<td>Magnifying desk lamp</td>
			<td>Brightech</td>
			<td>LightView Pro Flex 2</td>
			<td></td>
		</tr>
		<tr>
			<td>Metal Spatula</td>
			<td>N/A</td>
			<td>N/A</td>
			<td></td>
		</tr>
		<tr>
			<td>MgCl2 . 6H2O</td>
			<td>Fisher&#160;</td>
			<td>BP214-500</td>
			<td></td>
		</tr>
		<tr>
			<td>Micromanipulator (Right handed)</td>
			<td>WPI</td>
			<td>M3301R</td>
			<td></td>
		</tr>
		<tr>
			<td>Micromanipulator with digital display</td>
			<td>KOPF</td>
			<td>1940</td>
			<td></td>
		</tr>
		<tr>
			<td>Na2HPO4 . 7H2O</td>
			<td>Fisher&#160;</td>
			<td>S373-500</td>
			<td></td>
		</tr>
		<tr>
			<td>NaCl</td>
			<td>Sigma</td>
			<td>S7653-5Kg</td>
			<td></td>
		</tr>
		<tr>
			<td>NaH2PO4 . H2O</td>
			<td>Fisher&#160;</td>
			<td>S369-500</td>
			<td></td>
		</tr>
		<tr>
			<td>Neosporin</td>
			<td>Johnson &#38; Johnson</td>
			<td>N/A</td>
			<td>Apply topical oinment to prevent infection</td>
		</tr>
		<tr>
			<td>Parafilm</td>
			<td>Bemis</td>
			<td>DM-999</td>
			<td></td>
		</tr>
		<tr>
			<td>Rimadyl (Carprofen) 50mg/ml</td>
			<td>Zoetis</td>
			<td>N/A</td>
			<td>5 mg/kg, subcutaneous, for analgesia</td>
		</tr>
		<tr>
			<td>Scalpel</td>
			<td>N/A</td>
			<td>N/A</td>
			<td></td>
		</tr>
		<tr>
			<td>Stereotaxic allignment system</td>
			<td>KOPF</td>
			<td>1900</td>
			<td></td>
		</tr>
		<tr>
			<td>Sterile 27 gauge needle</td>
			<td>BD</td>
			<td>305109</td>
			<td></td>
		</tr>
		<tr>
			<td>Sterile cotton tip applicators (Solon)</td>
			<td>AMD Medicom</td>
			<td>56200</td>
			<td></td>
		</tr>
		<tr>
			<td>Sterile nylon sutures (5.0)</td>
			<td>Oasis</td>
			<td>MV-661</td>
			<td>Use non-absorable suture for closing the wound</td>
		</tr>
		<tr>
			<td>Sterile sharp scissors&#160;</td>
			<td>N/A</td>
			<td>N/A</td>
			<td></td>
		</tr>
		<tr>
			<td>Sterile surgical blades</td>
			<td>VWR</td>
			<td>55411-050</td>
			<td></td>
		</tr>
		<tr>
			<td>Surgical gloves (Nitrile)</td>
			<td>Ammex</td>
			<td>N/A</td>
			<td>Change gloves if there is suspision of contamination</td>
		</tr>
		<tr>
			<td>Tray</td>
			<td>N/A</td>
			<td>N/A</td>
			<td></td>
		</tr>
	</tbody>
</table>

osmotic minipump implantation for increasing glucose concentration in mouse cerebrospinal fluid

Diabetes increases the risk of cognitive decline and impairs brain function. Whether or not this relationship between high glucose and cognitive deficits is causal remains elusive. Moreover, whether these deficits are mediated by an increase in glucose levels in cerebrospinal fluid (CSF) and/or blood is also unclear. There are very few studies investigating the direct effects of high CSF glucose levels on central nervous system (CNS) function, especially on learning and memory, since current diabetes models are not sufficiently developed to address such research questions. This article describes a method to chronically increase CSF glucose levels for 4 weeks by continuously infusing glucose into the lateral ventricle using osmotic minipumps in mice. The protocol was validated by measuring glucose levels in CSF. This protocol increased CSF glucose levels to ~328 mg/dL after infusion of a 50% glucose solution at a 0.25 &#181;L/h flow rate, compared to a CSF glucose concentration of ~56 mg/dL in mice that received artificial cerebrospinal fluid (aCSF). Furthermore, this protocol did not affect blood glucose levels. Therefore, this method can be used to determine the direct effects of high CSF glucose on brain function or a specific neural pathway independently of changes in blood glucose levels. Overall, the approach described here will facilitate the development of animal models for testing the role of high CSF glucose in mediating features of Alzheimer&#39;s disease and/or other neurodegenerative disorders associated with diabetes.

Male mice were implanted with a cannula assembled to an osmotic minipump (Figure&#160;1) to chronically infuse aCSF or a 50% glucose solution into their lateral ventricles (Figure 2). CSF was collected 10 days after the surgery (Figure 3) to validate the efficacy of this procedure. The results showed an increase in the CSF glucose levels (mean: 327.7 mg/dL) in mice infused with 50% glucose compared to that (mean: 56.5 mg/dL) in mice...

Watch this Scientific Journal Video about Osmotic Minipump Implantation for Increasing Glucose Concentration in Mouse Cerebrospinal Fluid at JoVE.com

Osmotic Minipump Implantation for Increasing Glucose Concentration in Mouse Cerebrospinal Fluid

This article describes a detailed protocol to increase glucose concentration in the cerebrospinal fluid (CSF) of mice. This approach can be useful for studying the effects of high CSF glucose on neurodegeneration, cognition, and peripheral glucose metabolism in mice.

Diabetes increases the risk of cognitive decline and impairs brain function. Whether or not this relationship between high glucose and cognitive deficits ...

osmotic-minipump-implantation-for-increasing-glucose-concentration

Research

JoVE Journal

Biology

1.3K Views.  University of Rochester School of Medicine and Dentistry. This article describes a detailed protocol to increase glucose concentration in the cerebrospinal fluid (CSF) of mice. This approach can be useful for studying the effects of high CSF glucose on neurodegeneration, cognition, and peripheral glucose metabolism in mice.

Video: Osmotic Minipump Implantation for Increasing Glucose Concentration in Mouse Cerebrospinal Fluid

A Technique for Serial Collection of Cerebrospinal Fluid from the Cisterna Magna in Mouse 

Alzheimer's disease (AD) is a progressive neurodegenerative disease that is pathologically characterized by extracellular deposition of &#946;-amyloid peptide (A&#946;) and intraneuronal accumulation of hyperphosphorylated tau protein. Because cerebrospinal fluid (CSF) is in direct contact with the extracellular space of the brain, it provides a reflection of the biochemical changes in the brain in response to pathological processes. CSF from AD patients shows a decrease in the 42 amino-acid form of A&#946; (A&#946;42), and increases in total tau and hyperphosphorylated tau, though the mechanisms responsible for these changes are still not fully understood. Transgenic (Tg) mouse models of AD provide an excellent opportunity to investigate how and why A&#946; or tau levels in CSF change as the disease progresses. Here, we demonstrate a refined cisterna magna puncture technique for CSF sampling from the mouse. This extremely gentle sampling technique allows serial CSF samples to be obtained from the same mouse at 2-3 month intervals which greatly minimizes the confounding effect of between-mouse variability in A&#946; or tau levels, making it possible to detect subtle alterations over time. In combination with A&#946; and tau ELISA, this technique will be useful for studies designed to investigate the relationship between the levels of CSF A&#946;42 and tau, and their metabolism in the brain in AD mouse models. Studies in Tg mice could provide important validation as to the potential of CSF A&#946; or tau levels to be used as biological markers for monitoring disease progression, and to monitor the effect of therapeutic interventions. As the mice can be sacrificed and the brains can be examined for biochemical or histological changes, the mechanisms underlying the CSF changes can be better assessed. These data are likely to be informative for interpretation of human AD CSF changes.

A Technique for Serial Collection of Cerebrospinal Fluid from the Cisterna Magna in Mouse

Transgenic (Tg) mouse models of AD provide an excellent opportunity to investigate how and why A&#946; or tau levels in CSF change as the disease progresses in human patients. Here, we demonstrate a refined cisterna magna puncture technique for serial CSF sampling from the mouse.

Alzheimer's disease (AD) is a progressive neurodegenerative disease that is pathologically characterized by extracellular deposition of &#946;-amyloid ...

A Microfluidic Device for Quantifying Bacterial Chemotaxis in Stable Concentration Gradients

Chemotaxis allows bacteria to approach sources of attractant chemicals or to avoid sources of repellent chemicals. Bacteria constantly monitor the concentration of specific chemoeffectors by comparing the current concentration to the concentration detected a few seconds earlier. This comparison determines the net direction of movement. Although multiple, competing gradients often coexist in nature, conventional approaches for investigating bacterial chemotaxis are suboptimal for quantifying migration in response to concentration gradients of attractants and repellents. Here, we describe the development of a microfluidic chemotaxis model for presenting precise and stable concentration gradients of chemoeffectors to bacteria and quantitatively investigating their response to the applied gradient. The device is versatile in that concentration gradients of any desired absolute concentration and gradient strength can be easily generated by diffusive mixing. The device is demonstrated using the response of Escherichia coli RP437 to gradients of amino acids and nickel ions.

This protocol describes the development of a microfluidic device for investigating bacterial chemotaxis in stable concentration gradients of chemoeffectors.

Chemotaxis allows bacteria to approach sources of attractant chemicals or to avoid sources of repellent chemicals. Bacteria constantly monitor the ...

In vivo Imaging and Therapeutic Treatments in an Orthotopic Mouse Model of Ovarian Cancer

Human cancer and response to therapy is better represented in orthotopic animal models. This paper describes the development of an orthotopic mouse model of ovarian cancer, treatment of cancer via oral delivery of drugs, and monitoring of tumor cell behavior in response to drug treatment in real time using in vivo imaging system. In this orthotopic model, ovarian tumor cells expressing luciferase are applied topically by injecting them directly into the mouse bursa where each ovary is enclosed. Upon injection of D-luciferin, a substrate of firefly luciferase, luciferase-expressing cells generate bioluminescence signals. This signal is detected by the in vivo imaging system and allows for a non-invasive means of monitoring tumor growth, distribution, and regression in individual animals. Drug administration via oral gavage allows for a maximum dosing volume of 10 mL/kg body weight to be delivered directly to the stomach and closely resembles delivery of drugs in clinical treatments. Therefore, techniques described here, development of an orthotopic mouse model of ovarian cancer, oral delivery of drugs, and in vivo imaging, are useful for better understanding of human ovarian cancer and treatment and will improve targeting this disease.

In vivo Imaging and Therapeutic Treatments in an Orthotopic Mouse Model of Ovarian Cancer

Orthotopic animal models of ovarian cancer replicate better human disease and therefore enhance our understanding of cancer progression and tumor response to therapy. A mouse model receives an intrabursal injection of luciferase-expressing ovarian tumor cells. Treatment is administered via oral gavage. Tumor growth is monitored by in vivo imaging system.

Human cancer and response to therapy is better represented in orthotopic animal models.  This paper describes the development of an orthotopic mouse model ...

A 3D System for Culturing Human Articular Chondrocytes in Synovial Fluid

Cartilage destruction is a central pathological feature of osteoarthritis, a leading cause of disability in the US. Cartilage in the adult does not regenerate very efficiently in vivo; and as a result, osteoarthritis leads to irreversible cartilage loss and is accompanied by chronic pain and immobility 1,2. Cartilage tissue engineering offers promising potential to regenerate and restore tissue function. This technology typically involves seeding chondrocytes into natural or synthetic scaffolds and culturing the resulting 3D construct in a balanced medium over a period of time with a goal of engineering a biochemically and biomechanically mature tissue that can be transplanted into a defect site in vivo 3-6. Achieving an optimal condition for chondrocyte growth and matrix deposition is essential for the success of cartilage tissue engineering. 
 In the native joint cavity, cartilage at the articular surface of the bone is bathed in synovial fluid. This clear and viscous fluid provides nutrients to the avascular articular cartilage and contains growth factors, cytokines and enzymes that are important for chondrocyte metabolism 7,8. Furthermore, synovial fluid facilitates low-friction movement between cartilaginous surfaces mainly through secreting two key components, hyaluronan and lubricin 9 10. In contrast, tissue engineered cartilage is most often cultured in artificial media. While these media are likely able to provide more defined conditions for studying chondrocyte metabolism, synovial fluid most accurately reflects the natural environment of which articular chondrocytes reside in.
 Indeed, synovial fluid has the advantage of being easy to obtain and store, and can often be regularly replenished by the body. Several groups have supplemented the culture medium with synovial fluid in growing human, bovine, rabbit and dog chondrocytes, but mostly used only low levels of synovial fluid (below 20&#37;) 11-25. While chicken, horse and human chondrocytes have been cultured in the medium with higher percentage of synovial fluid, these culture systems were two-dimensional 26-28. Here we present our method of culturing human articular chondrocytes in a 3D system with a high percentage of synovial fluid (up to 100&#37;) over a period of 21 days. In doing so, we overcame a major hurdle presented by the high viscosity of the synovial fluid. This system provides the possibility of studying human chondrocytes in synovial fluid in a 3D setting, which can be further combined with two other important factors (oxygen tension and mechanical loading) 29,30 that constitute the natural environment for cartilage to mimic the natural milieu for cartilage growth. Furthermore, This system may also be used for assaying synovial fluid activity on chondrocytes and provide a platform for developing cartilage regeneration technologies and therapeutic options for arthritis.

A 3D system of culturing human articular chondrocytes in high levels of synovial fluid is described. Synovial fluid reflects the most natural microenvironment for articular cartilage, and can be easily obtained and stored. This system thus can be used for studying cartilage regeneration and for screening therapeutics for treating arthritis.

Cartilage destruction is a central pathological feature of osteoarthritis, a leading cause of disability in the US. Cartilage in the adult does not ...

Measuring the Osmotic Water Permeability Coefficient (Pf) of Spherical Cells: Isolated Plant Protoplasts as an Example

Studying AQP regulation mechanisms is crucial for the understanding of water relations at both the cellular and the whole plant levels. Presented here is a simple and very efficient method for the determination of the osmotic water permeability coefficient (Pf) in plant protoplasts, applicable in principle also to other spherical cells such as frog oocytes. The first step of the assay is the isolation of protoplasts from the plant tissue of interest by enzymatic digestion into a chamber with an appropriate isotonic solution. The second step consists of an osmotic challenge assay: protoplasts immobilized on the bottom of the chamber are submitted to a constant perfusion starting with an isotonic solution and followed by a hypotonic solution. The cell swelling is video recorded. In the third step, the images are processed offline to yield volume changes, and the time course of the volume changes is correlated with the time course of the change in osmolarity of the chamber perfusion medium, using a curve fitting procedure written in Matlab (the &#8216;PfFit&#8217;), to yield Pf.

Measuring the Osmotic Water Permeability Coefficient Pf of Spherical Cells: Isolated Plant Protoplasts as an Example

Measuring the osmotic water permeability coefficient (Pf) of cells can help understand the regulatory mechanisms of aquaporins (AQPs). Pf determination in spherical plant cell protoplasts presented here involves protoplasts isolation and numerical analysis of their initial rate of volume change as a result of an osmotic challenge during constant bath perfusion.

Studying AQP regulation mechanisms is crucial for the understanding of water relations at both the cellular and the whole plant levels. Presented here is ...

The Assembly and Application of 'Shear Rings': A Novel Endothelial Model for Orbital, Unidirectional and Periodic Fluid Flow and Shear Stress

Deviations from normal levels and patterns of vascular fluid shear play important roles in vascular physiology and pathophysiology by inducing adaptive as well as pathological changes in endothelial phenotype and gene expression. In particular, maladaptive effects of periodic, unidirectional flow induced shear stress can trigger a variety of effects on several vascular cell types, particularly endothelial cells. While by now endothelial cells from diverse anatomic origins have been cultured, in-depth analyses of their responses to fluid shear have been hampered by the relative complexity of shear models (e.g., parallel plate flow chamber, cone and plate flow model). While these all represent excellent approaches, such models are technically complicated and suffer from drawbacks including relatively lengthy and complex setup time, low surface areas, requirements for pumps and pressurization often requiring sealants and gaskets, creating challenges to both maintenance of sterility and an inability to run multiple experiments. However, if higher throughput models of flow and shear were available, greater progress on vascular endothelial shear responses, particularly periodic shear research at the molecular level, might be more rapidly advanced. Here, we describe the construction and use of shear rings: a novel, simple-to-assemble, and inexpensive tissue culture model with a relatively large surface area that easily allows for a high number of experimental replicates in unidirectional, periodic shear stress studies on endothelial cells.

Different levels and patterns of fluid shear are known to modulate endothelial gene expression, phenotype and susceptibility to disease. We discuss the assembly and use of 'shear rings': a model that produces unidirectional, periodic shear stress patterns. Shear rings are simple to assemble, economical and can produce high cell yields.

Deviations from normal levels and patterns of vascular fluid shear play important roles in vascular physiology and pathophysiology by inducing adaptive as ...

A Graphical User Interface for Software-assisted Tracking of Protein Concentration in Dynamic Cellular Protrusions

Filopodia are dynamic, finger-like cellular protrusions associated with migration and cell-cell communication. In order to better understand the complex signaling mechanisms underlying filopodial initiation, elongation and subsequent stabilization or retraction, it is crucial to determine the spatio-temporal protein activity in these dynamic structures. To analyze protein function in filopodia, we recently developed a semi-automated tracking algorithm that adapts to filopodial shape-changes, thus allowing parallel analysis of protrusion dynamics and relative protein concentration along the whole filopodial length. Here, we present a detailed step-by-step protocol for optimized cell handling, image acquisition and software analysis. We further provide instructions for the use of optional features during image analysis and data representation, as well as troubleshooting guidelines for all critical steps along the way. Finally, we also include a comparison of the described image analysis software with other programs available for filopodia quantification. Together, the presented protocol provides a framework for accurate analysis of protein dynamics in filopodial protrusions using image analysis software.

We present a software solution for semi-automated tracking of relative protein concentration along the length of dynamic cellular protrusions.

Filopodia are dynamic, finger-like cellular protrusions associated with migration and cell-cell communication. In order to better understand the complex ...

Trans-inner Cell Mass Injection of Embryonic Stem Cells Leads to Higher Chimerism Rates

In an effort to increase efficiency in the creation of genetically modified mice via ES Cell methodologies, we present an adaptation to the current blastocyst injection protocol. Here we report that a simple rotation of the embryo, and injection through Trans-Inner cell mass (TICM) increased the percentage of chimeric mice from 31% to 50%, with no additional equipment or further specialized training. 26 different inbred clones, and 35 total clones were injected over a period of 9 months. There was no significant difference in either pregnancy rate or recovery rate of embryos between traditional injection techniques and TICM. Therefore, without any major alteration in the injection process and a simple positioning of the blastocyst and injecting through the ICM, releasing the ES cells into the blastocoel cavity can potentially improve the quantity of chimeric production and subsequent germline transmission.

Here we present a protocol to increase chimera production without the use of new equipment. A simple orientation change of the embryo for injection can increase the number of embryos produced, and potentially reduce the timeline to germline transmission.

In an effort to increase efficiency in the creation of genetically modified mice via ES Cell methodologies, we present an adaptation to the current ...

In situ Hybridization for Sipunculus nudus Coelomic Fluid

In situ hybridization (ISH) is a very informative technique to present cellular distribution patterns of specific genes (e.g., mRNA and ncRNA) in tissues. The sipunculid worm Sipunculus nudus is a crucial fishery resource as it has high nutritional and medicinal values. Currently, the research on the molecular biology of Sipunculus nudus is still in its infancy. The purpose of this article is to develop a sensitive method for localizing specific mRNA in Sipunculus nudus coelomic fluid. The protocol includes detailed steps of ISH, including digoxigenin-labeled antisense and sense riboprobe preparation, coelomic fluid collection and section preparation, specific riboprobe hybridization, antibody incubation, coloration and post-coloration treatments. The representative results obtained from a successful experiment using this method are demonstrated. The protocol should be applicable to other Sipuncula species as well.

In situ Hybridization for Sipunculus nudus Coelomic Fluid

This protocol describes an effective in situ hybridization approach to detect the mRNA expression levels and spatial patterns of target genes in Sipunculus nudus coelomic fluid.

In situ hybridization (ISH) is a very informative technique to present cellular distribution patterns of specific genes (e.g., mRNA and ncRNA) in tissues. ...

Single Cell Collection of Trophoblast Cells in Peri-implantation Stage Human Embryos

Human implantation, the apposition and adhesion to the uterine surface epithelia and subsequent invasion of the blastocyst into the maternal decidua, is a critical yet enigmatic biological event that has been historically difficult to study due to technical and ethical limitations. Implantation is initiated by the development of the trophectoderm to early trophoblast and subsequent differentiation into distinct trophoblast sublineages. Aberrant early trophoblast differentiation may lead to implantation failure, placental pathologies, fetal abnormalities, and miscarriage. Recently, methods have been developed to allow human embryos to grow until day 13 post-fertilization in vitro in the absence of maternal tissues, a time-period that encompasses the implantation period in humans. This has given researchers the opportunity to investigate human implantation and recapitulate the dynamics of trophoblast differentiation during this critical period without confounding maternal influences and avoiding inherent obstacles to study early embryo differentiation events in vivo. To characterize different trophoblast sublineages during implantation, we have adopted existing two-dimensional (2D) extended culture methods and developed a procedure to enzymatically digest and isolate different types of trophoblast cells for downstream assays. Embryos cultured in 2D conditions have a relatively flattened morphology and may be suboptimal in modeling in vivo three-dimensional (3D) embryonic architectures. However, trophoblast differentiation seems to be less affected as demonstrated by anticipated morphology and gene expression changes over the course of extended culture. Different trophoblast sublineages, including cytotrophoblast, syncytiotrophoblast and migratory trophoblast can be separated by size, location, and temporal emergence, and used for further characterization or experimentation. Investigation of these early trophoblast cells may be instrumental in understanding human implantation, treating common placental pathologies, and mitigating the incidence of pregnancy loss.

Here, we describe a method for warming vitrified human blastocysts, culturing them through the implantation period in vitro, digesting them into single cells and collecting early trophoblast cells for further investigation.

Human implantation, the apposition and adhesion to the uterine surface epithelia and subsequent invasion of the blastocyst into the maternal decidua, is a ...