Name: Modeling Posthemorrhagic Hydrocephalus of Prematurity in Rats (Video) | JoVE Journal
Uploaded: 2025-03-28T12:35:00.000+00:00
Duration: 4 min 12 s
Description: Learn to model posthemorrhagic hydrocephalus of prematurity in neonatal rats using combined chorioamnionitis and intraventricular hemorrhage. This protocol replicates clinical features including macrocephaly and ventriculomegaly for translational research.

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5 minutes

The authors are grateful for the funding provided by the National Institutes of Health (R01HL139492), the Congressionally Directed Medical Research Program (W81XWH1810166, W81XWH1810167, W81XWH2210461, and W81XWH2210462), the Hydrocephalus Association, and the Rudi Schulte Research Institute.

The Animal Care and Use Committee (ACUC) at Johns Hopkins University approved all experimental procedures described herein. This protocol utilizes pregnant Sprague-Dawley rat dams and pups of both sexes.
1. Induction of chorioamnionitis on E18
NOTE: The in utero insult portion of this protocol has been previously published in detail, is summarized above, and is the subject of a separate JOVE protocol and video19,37,38,39,40,41,42,43,44,46. Briefly, pregnant female Sprague-Dawley rats undergo abdominal laparotomy on embryonic day 18 (E18) to induce chorioamnionitis, which includes TSHI and intraamniotic LPS administration.
<ol>
	<li>Anesthesia
	<ol>
		<li>Induce anesthesia in the E18 pregnant rat dam with 2-4% isoflurane.</li>
		<li>Remove the pregnant dam from the induction chamber and place the rat in the supine position on a draped surgical circulating water blanket set at 37 &#176;C.</li>
		<li>Apply ophthalmic ointment to prevent corneal drying. Gently squeeze a paw to confirm the absence of toe pinch reflex. Monitor the anesthetic depth every 15-20 min and increase the isoflurane in case of a positive toe pinch response.</li>
		<li>Administer buprenorphine extended release (0.1 mg/kg SC) at the nape of the neck.</li>
	</ol>
	</li>
	<li>Surgical preparation and scrub
	<ol>
		<li>Using standard sterile technique, shave the abdomen.</li>
		<li>Scrub the abdomen 3x with alternating betadine and 70% ethanol.</li>
		<li>Drape the animal using sterile surgical drapes.</li>
	</ol>
	</li>
	<li>Abdominal laparotomy
	<ol>
		<li>Make a 3 cm midline incision on the prepared abdominal skin with a scalpel.</li>
		<li>Use forceps and surgical scissors to hold up the abdominal fascial layer and make an incision of the avascular linea alba of the muscle layer to access the peritoneal cavity.</li>
		<li>Externalize the uterus.</li>
		<li>Isolate and clamp the uterine arteries with aneurysm clips for 60 min. Maintain the temperature and keep the intraabdominal contents moist with sterile saline.</li>
		<li>Remove the clips and inject 100 &#181;L of LPS (4 &#181;g/sac of LPS solution) into each amniotic sac of each fetus. Do not disturb the fetus or the placenta.</li>
		<li>Irrigate the uterine horns and field generously 3x with sterile saline.</li>
	</ol>
	</li>
	<li>Closing the laparotomy
	<ol>
		<li>Replace the uterine horns in the peritoneal cavity.</li>
		<li>Reapproximate the musculofascial layer edges and close using a running 3-0 suture.</li>
		<li>Reapproximate the skin layer and close the skin using a running 3-0 suture.</li>
		<li>Use a 26 G needle to subcutaneously inject 0.125% bupivacaine around the wound edges.</li>
		<li>For sham controls, perform the laparotomy for the same length of time to control for the duration of anesthesia. Do not clamp the arteries and do not administer any intraamniotic injections. At the conclusion of the procedure, close the laparotomy in two layers (abdominal muscle fascia and skin) using 3-0 suture. In all cases, the pups are born at term (E21/22) and are cared for by the dam.</li>
	</ol>
	</li>
</ol>
2. Preparation of lysed red blood cells on P1
<ol>
	<li>Collection of blood
	<ol>
		<li>Take one male and one female Sprague-Dawley rat pup at postnatal day 1 (P1) from a litter that experienced chorioamnionitis on E18. Rapidly decapitate each donor pup with dedicated surgical scissors. 
		NOTE: We use 1 male and 1 female pup for blood collection to eliminate a potential sex bias by representing each in the donor cohorts. Additionally, we use a sex-matched pair to guarantee sufficient volume and yield of lysed RBCs to inject their littermates. Typically, each donor pup yields enough lysed RBCs to perform ICV injection on a maximum of 4-5 littermates.</li>
		<li>Collect the blood immediately into a 2 mL microcentrifuge tube containing 0.2 mL of sterile saline, taking care to only collect free-flowing blood post decapitation and not scrape or squeeze to produce more blood as this leads to premature hemolysis. Vortex well. 
		NOTE: The exact amount of blood varies based on the individual donor animal and weight but should be maximum while maintaining the above precautions.</li>
		<li>Chop/mince blood clots with small surgical scissors.</li>
		<li>Centrifuge the blood suspension at 500 &#215; g for 10 min at 4 oC, remove the supernatant, and resuspend the pellet in 0.2 mL of sterile saline. Vortex well.</li>
		<li>Chop/mince post vortex residual blood clots with small surgical scissors.</li>
		<li>Repeat steps 2.1.4-2.1.5 twice more for a total of 3x, cleaning surgical scissors with 70% ethanol spray in between each round of lysing.</li>
	</ol>
	</li>
	<li>Lysis of red blood cells
	<ol>
		<li>After the final centrifugation, add 0.25 mL of sterile saline to the pellet; vortex well.</li>
		<li>Place the suspension on dry ice for 5 min.</li>
		<li>Remove the suspension from dry ice, place it in an incubator set at 37.5 oC for 5 min until completely thawed, and vortex well.</li>
		<li>Repeat the freeze and thaw cycles for a total of 3x (three freezes and three thaws).</li>
		<li>At the conclusion of the last thaw, vortex and perform a quick spin. The RBCs are now lysed and ready to use. 
		NOTE: The mixture should be an opaque tomato juice-like color and be easily drawn into syringes.</li>
	</ol>
	</li>
</ol>
3. Intracerebroventricular injections of lysed red blood cells on P1
<ol>
	<li>Anesthesia using hypothermia
	<ol>
		<li>Place a small platform on wet ice to cool.</li>
		<li>Place a dry laboratory wipe on top to protect the pup&#8217;s skin. 
		NOTE: This flat cold surface is used for anesthetizing and injecting the pups.</li>
		<li>Transfer on pup (aged P1) from the warming pad onto the task wipe atop cold platform to induce anesthesia by hypothermia.</li>
		<li>Confirm the depth of anesthesia by squeezing a paw and confirming the absence of the toe pinch reflex.</li>
		<li>Set an external surgical lamp to its brightest settings.</li>
		<li>With an assistant using their index and middle finger to gently maintain the animal&#8217;s head midline, transilluminate the skull to visualize the lateral ventricles through the skull. Identify bregma by visualizing the superior sagittal sinus (midline) through the skin and palpation of the coronal suture with fine forceps as intersecting landmarks.</li>
	</ol>
	</li>
	<li>ICV injection
	<ol>
		<li>Wipe the head of the anesthetized pup with a cotton swab soaked in 70% ethanol.</li>
		<li>Identify and mark the injection site as 1 mm lateral from the sagittal suture, halfway between lambda and bregma.</li>
		<li>After visualization, use a 0.3 mL, 8 mm long, 31 G insulin syringe with an ultrafine percutaneous needle to inject 20 &#181;L of lysed RBCs into the right lateral ventricle. Insert the needle straight downward using freehand technique to a depth of approximately half the needle length and inject and remove the needle slowly (injection and removal process over approximately 10-15 s).</li>
		<li>Leave the needle in place for several seconds after the injection to prevent egress of the injected lysed RBCs.</li>
		<li>Repeat with the left lateral ventricle and inject 20 &#181;L of lysed RBCs.</li>
		<li>Place the pup on a warming pad set at 37.5 oC to recover from anesthesia.</li>
		<li>Record the sex of the pup and assign a unique animal identifier.</li>
		<li>Return the pup to the home cage only after full recovery on the warming pad and regaining of consciousness such that animal can safely maintain sternal recumbency.</li>
		<li>Monitor all rat pups daily for health and wellness.</li>
	</ol>
	</li>
</ol>
4. Confirmation of successful bilateral intraventricular hemorrhage on P2
<ol>
	<li>Head ultrasound
	<ol>
		<li>To prepare for head ultrasound, remove the P2 pups from their home cage.</li>
		<li>Place ultrasound gel on the ultrasound probe and position the probe over the top of the cranium.</li>
		<li>With extremely light pressure, move the probe to visualize the ventricles. Confirm bilateral hyperechogenicity in the lateral ventricles representing IVH.</li>
	</ol>
	</li>
</ol>
5. Confirmation of successful posthemorrhagic hydrocephalus
<ol>
	<li>Measurement of intraaural distance (IAD), a surrogate for head circumference, to confirm macrocephaly
	<ol>
		<li>To prepare for measurement, acquire a small tape measure appropriate for measuring head circumference, ideally with clearly visualized millimeter designations.</li>
		<li>Have a masked observer remove the pup from their home cage.</li>
		<li>While gently holding the pup, measure the distance from ear to ear (intraaural distance, IAD) and record the value in millimeters.</li>
		<li>Repeat IAD daily from P1 to P15 and graph the values. Serially track IAD and measure again at P21 upon relocating pups to new cages physically separated from the dam (which is the standard time point for pup weaning). Repeat IAD subsequently every 5 days starting at P25 until P60.</li>
	</ol>
	</li>
	<li>Measurement of opening pressure to confirm elevated intracranial pressure
	<ol>
		<li>Have a masked observer remove the pup from its home cage.</li>
		<li>Anesthetize with 75-100 mg/kg intraperitoneal (IP) ketamine and 5-10 mg/kg IP xylazine in preparation for euthanasia.</li>
		<li>Check the depth of anesthesia by squeezing a paw and confirming the absence of the toe pinch/pedal reflex.</li>
		<li>Insert a small needle (31 G) connected to a manometer into the cervicomedullary junction CSF space.</li>
		<li>Record the opening pressure on the manometer.</li>
		<li>Remove the needle and decapitate the rat with sharp scissors and proceed with tissue collection.</li>
	</ol>
	</li>
	<li>Ex-vivo magnetic resonance imaging (MRI) for assessment of ventriculomegaly
	<ol>
		<li>Anesthetize with 75-100 mg/kg intraperitoneal (IP) Ketamine and 5-10 mg/kg IP Xylazine in preparation for euthanasia.</li>
		<li>Check the depth of anesthesia by squeezing a paw and confirming the absence of the toe pinch/pedal reflex.</li>
		<li>Perfuse the rats with phosphate-buffered saline (PBS), followed by 4% paraformaldehyde (PFA) until fixed well.</li>
		<li>Remove the brain and drop-fix the brain in 4% PFA</li>
		<li>Embed the brain in 2% agarose in a 50 mL conical tube. Let it stand at room temperature.</li>
		<li>Transfer the brain to the MRI scanner for ex vivo MRI.</li>
		<li>Run 11.7T MRI as follows: T2 Turbo RARE; TE/TR = 30.0/3000 ms; avg = 2; Echo spacing = 10.000 ms; RARE factor = 8; number of slices = 30; slice thickness = 1 mm; image size = 128 x 128; FOV = 28 mm x 28 mm; slice resolution = 0.219 x 0.219 mm2; FA = 90.0&#176;. 
		NOTE: While MRI imaging provides evidence of successful PHHP modeling, it is not required to scan all brains in a given cohort&#160;to verify the PHHP. IAM and ICP measurement is sufficient to verify as described above. Ultimately, the ability of an investigator to perform in vivo or ex vivo MRI will be dependent on a variety of factors such as MRI scanner access, funds, and technical ability. This step is particularly useful for validation when newly incorporating the PHHP model. It is key to note that in the absence of documented signs of increased intracranial pressure, such as elevated opening pressure, isolated findings of ventriculomegaly on MRI imaging does not represent hydrocephalus.</li>
	</ol>
	</li>
</ol>

The authors have no conflicts of interest to disclose.

This protocol for the induction of PHHP allows for rigorous, quantifiable, and clinically translatable outcome measures of brain structure and function concomitant with phenotypic hallmarks of hydrocephalus, including chronic elevation of ICP, ventriculomegaly, and macrocephaly, from birth to adulthood4. Biochemical, histological, and functional assays can be used to evaluate the health of the choroid plexus, ependyma, and glymphatic system, as well as gray and white matter19</su...

Posthemorrhagic hydrocephalus of prematurity (PHHP) remains a substantial public health concern. Defined by symptomatic accumulation of cerebrospinal fluid (CSF) concomitant with elevated intracranial pressure (ICP) secondary to intraventricular hemorrhage (IVH), PHHP is a severe manifestation of encephalopathy of prematurity and a significant contributor to the global burden of prematurity and acquired hydrocephalus1,2. Globally, approximately 400,000 infants each year are born with or acquire the lifelong burden of hydrocephalus3 and many die due to lack of treatment3. PHHP is common in developed countries in very preterm infants (&#60;32 weeks&#39; gestation) with severe IVH, and often affects the sickest of infants who are already suffering from other life-threatening co-morbidities4,5.
The only available treatment for hydrocephalus is surgery6. Surgical procedures yield better longevity when infants are older than 6 months at the time of the first permanent intervention, whether for a ventriculoperitoneal (VP) shunt to divert cerebrospinal fluid (CSF), endoscopic third ventriculostomy (ETV), or ETV with choroid plexus coagulation (ETV-CPC)7. The most common option, VP shunts, often fail within a year and predispose children to a lifetime of complications, repeat surgeries, and hospitalizations at a tremendous cost to the child, the family, and society.8 In particular, the anxiety from a shunt potentially failing at any time is burdensome to families9. Care for children with symptomatic hydrocephalus, including frequent surgeries, is a leading cause of pediatric healthcare expenditures10,11,12,13,14. The annual estimated cost for shunt-related expenditures in children was $2 billion in 200315. While children with shunts comprise only 0.6% of hospital admissions, they generate 3.1% of pediatric hospital charges15. Thus, the discovery of safe, non-surgical therapies for the treatment of PHHP is paramount.
In infants, PHHP develops after IVH over a clinical time course that lasts weeks to months after the initial identification of the brain bleed. A study conducted by the Hydrocephalus Clinical Research Network (HCRN) confirmed that VP shunts remain the best surgical option for neonates with PHHP16. Even for children with PHHP in high-income countries with access to skilled pediatric neurosurgical care, outcomes are far from optimal, with &#62;50% of shunts placed in infants with PHHP requiring surgical revision within the first 2 years8. Despite the clear need to identify safer, more effective treatments for PHHP, research has faced obstacles. Progress has been hampered in part because the preclinical literature on PHHP often fails to appropriately distinguish ventriculomegaly caused by hydrocephalus ex vacuo17,18&#160;from&#160;symptomatic hydrocephalus with macrocephaly19,20. Indeed, developmental models of hydrocephalus should include progressive macrocephaly and/or measurements of elevated ICP1.
Merging clinical and preclinical insights has improved study design and propelled our understanding of PHHP2. Studies conducted in diverse centers throughout the globe have shown that IVH is most common in very preterm neonates secondary to chorioamnionitis21,22,23,24,25,26,27,28. In addition to placental infection and inflammation, neonatal sepsis is an additional important risk factor and can play a central role in the progression from IVH to ventriculomegaly to symptomatic PHHP and subsequent surgical intervention29. Preclinical and clinical data support that blood-borne inflammation can cause hydrocephalus20, and systemic inflammation increases secretion of CSF by the choroid plexus30. Further, adults with subarachnoid hemorrhage and IVH who also suffer from sepsis are much more likely to require a shunt31. More recent literature has confirmed that inflammation reduces ependymal motile cilia propulsion of CSF19,20,32 and CSF reabsorption by the glymphatic system33,34,35,36. Overall, systemic inflammation is a key pathophysiological and clinical driver in PHHP1.
Considering these findings, we created an age-appropriate preclinical model of PHHP. This model combines IVH in the immediate and early postnatal period with chorioamnionitis, the principal cause of preterm birth19. This experimental approach begins in utero, with the placental insufficiency, placental inflammation, and intraamniotic inflammation that defines chorioamnionitis7,8,9,10,11,12,13,14,15,16,17,18,19,20,21,22, 
23,24,25,26,27,28,29,30,31,32,33,34,35,36,37,38,39,40,41,42, 
43,44,45. Specifically, we recapitulate a fetal inflammatory response syndrome, placental neutrophilia, and proinflammatory CNS microenvironment in the preterm period via abdominal laparotomy in pregnant rat dams on embryonic day 18 (E18)37,38,39,40,41,42,43,44,45. Intrauterine injury is induced by temporary bilateral uterine artery occlusion leading to transient systemic hypoxia-ischemia (TSHI) followed by intraamniotic injection of lipopolysaccharide (LPS)37,38,39,40,41,42,43,44,45. Subsequently, to perturb CSF dynamics and catalyze the development of hydrocephalus in the live-born pups, IVH is induced on postnatal day 1. This is accomplished with bilateral intracerebroventricular injection (ICV) of littermate lysed red blood cells (RBCs) into the lateral ventricles19,37,44. Pups are then studied as hydrocephalus develops and throughout their lifespan.

<table><tbody><tr><td>70% ethanol&amp;nbsp;</td><td>Pharmco&amp;nbsp;</td><td>111000200</td><td>Diluted to 70%&amp;nbsp;</td></tr><tr><td>Betadine surgical scrub</td><td>Cardinal Health</td><td>NDC-67618-151-17</td><td></td></tr><tr><td>Blunt Forceps</td><td>Roboz</td><td>RS-8100</td><td></td></tr><tr><td>Bravmini Plus Cordless Rechargeable Trimmer&amp;nbsp;</td><td>Wahl&amp;nbsp;</td><td>41590-0438</td><td></td></tr><tr><td>Carbon Steel Surgical blades&amp;nbsp;</td><td>Bard-Parker</td><td>371151-11</td><td></td></tr><tr><td>centrifuge&amp;nbsp;</td><td>Eppendorf</td><td>5424R</td><td></td></tr><tr><td>Cotton Gauze Sponge</td><td>Fisherbrand</td><td>22-362-178</td><td>Small, 6 inch sterile</td></tr><tr><td>Cotton-tipped Applicators</td><td>Fisherbrand</td><td>23-400-114</td><td>30 G 1</td></tr><tr><td>Eye Lubricant</td><td>Refresh Lacri Lube</td><td>75929</td><td></td></tr><tr><td>Far infrared warming pad</td><td>Kent scientific</td><td>RT-0501&amp;nbsp;</td><td></td></tr><tr><td>Incubator -&amp;nbsp; Genie Temp-Shaker 100&amp;nbsp;</td><td>Scientific Industries</td><td>SI-G100</td><td></td></tr><tr><td>Insulin Syringes</td><td>BD</td><td>328438</td><td>0.3 cc 3 mm 31 G, ultrafine&amp;nbsp;</td></tr><tr><td>Isoflurane</td><td>Covetrus&amp;nbsp;</td><td>11695067772&amp;nbsp;</td><td></td></tr><tr><td>Ketamine hydrochloride injection</td><td>Dechra&amp;nbsp;</td><td>17033-101-10</td><td></td></tr><tr><td>Kimwipes</td><td>Kimtech Science</td><td>BXTNI141300</td><td></td></tr><tr><td>LPS 011B4</td><td>Sigma</td><td>L2630</td><td></td></tr><tr><td>microcentrifuge tubes</td><td>Thermo Fisher Scientific</td><td>3453</td><td>2.0 mL</td></tr><tr><td>Needle</td><td>BD</td><td>305122</td><td>1 mL</td></tr><tr><td>Needle</td><td>BD</td><td>305128</td><td>25 G 5/8</td></tr><tr><td>Needle Holders</td><td>Kent Scientific Corp.</td><td>INS14109</td><td>12.5 CM STR</td></tr><tr><td>OR Towels</td><td>Cardinal Health</td><td>287000-008</td><td></td></tr><tr><td>Paper measuring tape</td><td>Cardinal Health</td><td>SKU &amp;nbsp;</td><td></td></tr><tr><td>Saline Solution, 0.9%</td><td>Sigma</td><td>S8776</td><td></td></tr><tr><td>Scissors</td><td>Roboz</td><td>RS-6808</td><td></td></tr><tr><td>SomnoSuite</td><td>Kent Scientific</td><td>SS6823B&amp;nbsp;</td><td></td></tr><tr><td>Sterile Alcohol Prep Pads</td><td>Fisherbrand</td><td>06-669-62</td><td>Sterile</td></tr><tr><td>Surgical gloves</td><td>Biogel</td><td>40870</td><td></td></tr><tr><td>Surgical Scissors</td><td>Roboz</td><td>RS-5880</td><td></td></tr><tr><td>Surgical Scissors</td><td>EST</td><td>14002-16</td><td></td></tr><tr><td>Syringe</td><td>BD</td><td>309628</td><td></td></tr><tr><td>T/Pump (Heat Therapy Pump)</td><td>Stryker Medical&amp;nbsp;</td><td>TP700</td><td></td></tr><tr><td>Vessel Clips</td><td>Kent Scientific Corp.</td><td>INS14120</td><td>30 G Pressure</td></tr><tr><td>Xylazine injection&amp;nbsp;</td><td>vet one&amp;nbsp;</td><td>NDC 13985-704-10</td><td></td></tr></tbody></table>

modeling posthemorrhagic hydrocephalus of prematurity in rats

Modeling Posthemorrhagic Hydrocephalus of Prematurity in Rats

Posthemorrhagic hydrocephalus of prematurity (PHHP) is a serious sequela of severe intraventricular hemorrhage (IVH) in very preterm infants less than 32 weeks gestational age (GA). PHHP is defined by the accumulation of cerebrospinal fluid (CSF) associated with clinical symptoms of elevated intracranial pressure (ICP). Infants with PHHP suffer lifelong shunt dependence, with half requiring repeat surgery in the first year of life and many requiring multiple additional surgeries throughout the lifespan. Prenatal chorioamnionitis predisposes preterm infants to severe IVH and the need for surgical treatment of PHHP trends with neonatal sepsis. These clinical features suggest that systemic inflammation is an integral component of PHHP pathophysiology. 
Here, we define an animal model that recapitulates all clinical aspects and essential features of PHHP in rats. The goal of this protocol is to illustrate how in utero chorioamnionitis and postnatal IVH using lysed red blood cells can be combined to yield PHHP. This preclinical approach yields progressive macrocephaly and domed craniums, elevated intracranial pressure, and ventriculomegaly that can be detected via magnetic resonance imaging (MRI) or via microscopy. In addition to sustained disruption in CSF dynamics, rats also have cognitive delay and functional disability into adulthood. Accordingly, this preclinical platform facilitates unique and unparalleled translational studies of PHHP that can incorporate molecular, cellular, biochemical, histologic, imaging, and functional outcome measures. It can also be used for rigorous analysis of the choroid plexus, ependymal motile cilia, and glymphatic system in parallel. Last, it can also be an invaluable preclinical tool for the investigation of novel surgical intervention strategies and non-surgical therapeutic approaches for the treatment of hydrocephalus.

Using this model, hydrocephalus develops in the days and weeks after injection of lysed red blood cells. A representation of a typical experimental design and progression of hydrocephalus is provided in Figure 1. We evaluated 5-6 sham animals and 6-8 PHHP animals per group. As juveniles, rats with PHHP exhibited macrocephaly (Figure 2), elevated intracranial pressure (Figure 3), and ventriculomegaly (Figure 4</s...

Posthemorrhagic hydrocephalus of prematurity (PHHP) can be modeled in neonatal rats by combining chorioamnionitis and intraventricular hemorrhage. The combination of these prenatal and postnatal events accurately recapitulates the clinical hallmarks of PHHP, including macrocephaly, ventriculomegaly, and elevated intracranial pressure, through the lifespan.

Posthemorrhagic hydrocephalus of prematurity (PHHP) is a serious sequela of severe intraventricular hemorrhage (IVH) in very preterm infants less than 32 ...

Our laboratory seeks to characterize cellular and molecular pathophysiologic mechanisms of different perinatal brain injuries, with a specific interest being in posthemorrhagic hydrocephalus of prematurity, or PHHP. Our goal is to uncover novel, translatable therapies for these challenging diseases. This protocol helps us better understand the damage done to the developing brain after intraventricular hemorrhage and hydrocephalus.With this model, we are uncovering potential strategies to reverse that brain damage and develop new and less invasive treatments. This protocol recapitulates essential clinical features of PHHP in rats by combining in utero chorioamnionitis and postnatal IVH using lysed red blood cells. The protocol yields rats with sustained macrocephaly, ventriculomegaly, increased intracranial pressure, and functional disability into adulthood, facilitating translational PHHP studies with a full spectrum of outcome measures.To begin, obtain blood from a male and a female Sprague-Dawley rat pup at postnatal day one from a litter that experienced in utero chorioamnionitis on embryonic day 18. Vortex the tube well to mix the contents. Using small surgical scissors, chop and mince any blood clots in the tube.Centrifuge the blood suspension at 500g for 10 minutes at four degrees Celsius. Remove the supernatant. Resuspend the pellet in 0.2 milliliters of sterile saline, and vortex well to ensure proper mixing.Chop and mince any residual blood clots in the suspension post-vortex using small surgical scissors. After the final centrifugation, add 0.25 milliliters of sterile saline to the pellet and vortex the tube well to mix the contents. Place the suspension on dry ice for five minutes.Now transfer the suspension into an incubator set at 37.5 degrees Celsius for five minutes until completely thawed, and vortex the suspension well after thawing. Repeat the freeze and thaw cycles three times in total. After the final thaw, vortex the tube and perform a quick spin in the centrifuge.Position a small platform on wet ice and place a dry laboratory wipe on the platform to protect the pup's skin during the procedure. After anesthetizing the pup by hypothermia, position an external surgical lamp set to its brightest settings. Transilluminate the skull to visualize the lateral ventricles through the skull.Use a 0.3-milliliter, eight-millimeter long, 31-gauge insulin syringe with an ultra-fine percutaneous needle to inject 20 microliters of lysed red blood cells into the right lateral ventricle. Rats with posthemorrhagic hydrocephalus of prematurity, or PHHP, exhibited enlarged, domed craniums and macrocephaly, as shown by an increased head circumference surrogate measure observable as juveniles. Rats with PHHP had increased intra-aural distance and increased intracranial pressure on postnatal day 21 compared to sham controls.Ventriculomegaly and increased ventricular volume in rats with PHHP were visible through MRI, confirmed by comparison with sham controls.

modeling-posthemorrhagic-hydrocephalus-of-prematurity-in-rats

Research

JoVE Journal

Neuroscience

The postnatal rat model for hypoxic-ischemic brain injury is a well-established model of human neonatal hypoxic ischemic encephalopathy (HIE). In this article, we describe the model of HIE in post-natal rat pups.

The Hypoxic Ischemic Encephalopathy Model of Perinatal Ischemia

We established mouse models of periventricular leukomalacia (PVL), the predominant brain injury in premature infants characterized by periventricular white matter lesions. Hypoxia/ischemia with/without systemic infection are the primary causes of PVL. Unilateral carotid ligation and hypoxia exposure with/without lipopolysaccharide injection creates PVL-like lesions in P6 mice.

Mouse Models of Periventricular Leukomalacia

Aneurysmal subarachnoid hemorrhage (SAH) is bleeding that occurs into the subarachnoid space when an aneurysm ruptures. While the morbidity and mortality from this event has been on a decline due to improved treatment approaches, the risk of vasospasm after subarachnoid hemorrhage continues to be the same as it was several years ago. The importance of establishing a comprehensive and reproducible animal model to identify initiating events of cerebral vasospasm has been the focus of research...

A Low Mortality Rat Model to Assess Delayed Cerebral Vasospasm After Experimental Subarachnoid Hemorrhage

This video demonstrates a method to generate a rat model of neonatal intraventricular hemorrhage. In this procedure, an anesthetized rat pup is injected with hemoglobin into the lateral ventricle of the brain. The hemoglobin causes oxidative stress and releases heme, simulating intraventricular hemorrhage. The resulting damage to brain tissues, driven by reactive oxygen species and inflammatory cytokines, leads to ventricular enlargement, a common consequence of intraventricular hemorrhage.

Modeling Neonatal Intraventricular Hemorrhage Through Intraventricular Injection of Hemoglobin

Source: Jantzie, L. L., et. al. Modeling Encephalopathy of Prematurity Using Prenatal Hypoxia-ischemia with Intra-amniotic Lipopolysaccharide in Rats. J. Vis. Exp. (2015)This video demonstrates a protocol to induce hypoxic-ischemic and inflammatory conditions in fetal rats by restricting uterine blood flow and injecting LPS, modeling neuroinflammation and disrupted brain development relevant to preterm birth studies.

Inducing Hypoxic-Ischemic and Inflammatory Conditions to Model Encephalopathy in Prenatal Rats

Encephalopathy of prematurity encompasses the central nervous system abnormalities associated with injury from preterm birth. This report describes a clinically relevant rat model of in utero transient systemic hypoxia-ischemia and intra-amniotic lipopolysaccharide administration (LPS) that mimics chorioamnionitis, and the related impact of infectious stimuli and placental underperfusion on CNS development.

Modeling Encephalopathy of Prematurity Using Prenatal Hypoxia-ischemia with Intra-amniotic Lipopolysaccharide in Rats

Mild intrauterine hypoperfusion was produced by artery stenosis with metal microcoils wrapped around the uterine and ovarian arteries in rats at embryonic day 17. This procedure produced prenatal hypoperfusion and intrauterine growth restriction.

A Rat Model of Mild Intrauterine Hypoperfusion with Microcoil Stenosis

Monitoring intracranial pressure in rodent models of nontraumatic intraventricular hemorrhage is not common in the current literature. Herein, we demonstrate a technique for measuring intracranial pressure, mean arterial pressure, and cerebral perfusion pressure during intraventricular hemorrhage in a rat animal model.

Intracranial Pressure Monitoring In Nontraumatic Intraventricular Hemorrhage Rodent Model

We present a model of neonatal intraventricular hemorrhage using rat pups that mimics the pathology seen in humans.

Hemorrhagic shock kills 1.9 million people worldwide every year. Small animals are frequently used as hemorrhagic shock models but are associated with issues of standardization, reproducibility, and clinical significance, thus limiting their relevance. This article describes developing a new clinically relevant hemorrhagic shock model in rats.

Developing a Clinically Relevant Hemorrhagic Shock Model in Rats

Learn to model posthemorrhagic hydrocephalus of prematurity in neonatal rats using combined chorioamnionitis and intraventricular hemorrhage. This protocol replicates clinical features including macrocephaly and ventriculomegaly for translational research.

Modeling Posthemorrhagic Hydrocephalus of Prematurity in Rats

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