Chapter 1 - Springer Link

Chapter 1 A Method to the Madness: Producing the Neonatal Ventral Hippocampal Lesion Rat Model of Schizophrenia R. Andrew Chambers and Barbara K. Lipska Abstract The neonatal ventral hippocampal lesion (NVHL) rat model of schizophrenia has demonstrated broad heuristic utility as an investigative platform encompassing many of the behavioral, neurobiological, and developmental aspects of this devastating neuropsychiatric illness affecting 1% of all human beings. This chapter serves as an essential description of materials and methods for generating and verifying the NVHL model in rats, which continues to hold significant potential in helping us understand schizophrenia, comorbid disorders, and their neurodevelopmental dynamics. Many of the approaches described here can be modified or adapted for producing other types of neurodevelopmental models of behavioral disorders. Key words: Ventral hippocampus, neonatal lesions, ibotenic acid, neurodevelopmental, methods.

1. Introduction The neonatal ventral hippocampal lesion (NVHL) rat model, developed at St. Elizabeth’s Hospital in Washington, DC, was first described in 1993 from the intramural program of the NIMH (1). It has become one of the most intensively studied animal models of schizophrenia ever developed, being the topic of over 120 primary research publications by at least a dozen independent research groups worldwide (2). Aside from representing a major advance in schizophrenia research in terms of providing the first significant non-pharmacological model of schizophrenia (i.e., not produced by exposure to psychostimulant or hallucinogenic drugs), this model includes multiple features that make it an attractive investigative platform. As a model of a complex neuropsychiatric syndrome that encompasses many neurobiological, P. O’Donnell (ed.), Animal Models of Schizophrenia and Related Disorders, Neuromethods 59, DOI 10.1007/978-1-61779-157-4_1, © Springer Science+Business Media, LLC 2011

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endophenotypic, and clinical/diagnostic features, which might seem etiologically unrelated, it succeeds in reproducing most of these phenomena in detail, if not in broad strokes, as a result of one experimental intervention: developmental damage of the ventral hippocampus in post-natal rats (see (2) for a comprehensive review). So, for example, instead of representing a paradigm that can test rats for a single schizophrenic trait, such as deficits in pre-pulse inhibition (PPI) of auditory stimuli, the model itself produces deficits in PPI, and many other schizophrenialike features including deficits in latent inhibition (LI), positivelike symptoms (e.g., behavioral hyper-responsiveness to novelty, stress, and psychostimulants that are reducible with neuroleptic treatment), cognitive symptoms (e.g., spatial working memory deficits), and neuroleptic-unresponsive negative symptoms (e.g., abnormalities in grooming and social behavior). Neurobiologically, NVHLs produce a host of structural and functional alterations that are downstream in anatomical space and developmental time from the initial lesion and that also mimic neuroimaging or post-mortem findings in human schizophrenia. Timely to growing interest in the periadolescent neurodevelopmental basis of schizophrenia and many other psychiatric disorders, the positive symptom-like behavioral features of the model show periadolescent emergence, in parallel with progressively abnormal biological or physiological measures of prefrontal cortical dysfunction (3). Moreover, the distributed neurocircuit effects of the lesion also represent mechanisms informative to understanding complex co-morbidities spanning mental illnesses such as addictions (4). Finally, the model itself or many of the techniques for producing it may be used to develop more specific experimental interventions in modeling a wide variety of other mental disorders involving alternative developmental time points, anatomical sites, or bioactive agents of lesion delivery, or combinations of lesions with particular environmental or genetic backgrounds. Successfully producing NVHL rats in sufficient quantities, while similar to approaches commonly used in stereotaxic brain interventions in adult rats, requires more of a nuanced, skilled touch and several special modifications. For the second author, originally developing this model accurately (e.g., achieving >80% lesion accuracy) from scratch required the sacrifice of well over 1000 rat pups, and working though numerous “wrong directions.” Even with the bugs worked out, and as trained directly from the inventor herself, the first author required nearly 4 months of effort and >100 pups to gain >66% proficiency. It is our goal to provide this chapter as a fairly complete and definitive resource for other investigators who are starting out in producing NVHL or similar developmental models so that they may get up and running with sufficient number of rats more quickly. We describe the essential materials and methods for producing the

Neonatal Hippocampal Lesion Model

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NVHL lesion and verifying its accuracy in adulthood. Section 4 describes trouble areas, pitfalls, and miscellaneous advice in this experimental approach.

2. Materials 2.1. Rats

Any rat strain of either gender may be used with the NVHL model, but the majority of papers (>90%) have examined Sprague–Dawleys and probably at least three-quarters of papers have focused on males. Thus if an experiment aims to reference or contextualize experimental results to the bulk of the literature, the best bet is to use Sprague–Dawley males. Alternatively, to explore gender or genetic biases with the lesion, both genders or Spragues with another strain are used in single experiments. The lesions are made in (and stereotaxic coordinates are designed for) 7-day-old rat pups (PD-7). This developmental point was chosen to approximate the second to third trimester equivalent of human fetal brain development, when the hippocampus is undergoing a crucially sensitive period of network formation, thought to make it vulnerable to a host of infectious, hormonal, neuroimmunological, or anoxic risk factors implicated in the early pathogenesis of human schizophrenia. Given this background, and the potential effects of other sources of perinatal stress, it is essential that rats are born in the laboratory from pregnant dams, either bred in or arriving in the laboratory housing facility no more than 17 days into gestation (i.e., 4 or more days before parturition on gestational day 21). Orders for pregnant dams at 13–17 days of gestation usually require 2–4 weeks advanced notice. In our experience, a given pregnant dam will typically provide 4–6 male rats/litter of appropriate weight range for the surgery. With one stereotaxic setup, and performing lesions proficiently at full speed, surgeries may proceed at about 4 rats/h. This rate can be increased with more stereotaxic setups (e.g., 6–8 rats/h with two setups). Depending on the expected rate of surgery, pregnant dams should be ordered with gestational due dates 1–2 days apart to avoid traffic jams of pups available for surgery.

2.2. Ibotenic Acid and Artificial CSF

Ibotenic acid (α-amino-3-hydroxy-5-isoxazoleacetic acid) is an NMDA receptor agonist, naturally produced by the Amanita mushroom species. In low doses delivered generally to the CNS, it has psychedelic effects, and interestingly, a portion of the compound is metabolized to muscimol, a potent and selective GABAA receptor agonist. In concentrated doses delivered to specific brain regions, it causes lethal neuronal calcium influx via the NMDA

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receptor. However, as an advantage among lesion techniques, it predominantly kills cells with cell bodies in the vicinity of the high dose and leaves axonal projections through the “blast zone” intact. Ibotenic acid (IBO) is available in 5 mg quantities (e.g., from Sigma) (>$350) providing enough toxin, theoretically, to lesion up to 800 rats. Ibotenic acid for neonatal lesioning is dissolved in artificial CSF (aCSF) vehicle, which after mixing and storing in Eppendorf tubes may be kept in –80◦ C freezer for >6 months and possibly up to 1 year without losing effectiveness. At room temperature and subjected to ambient light, IBO will lose potency within 6 h. The aCSF is a solution of electrolytes at physiologic pH that is inert when administered intrathecally to the rat. Beyond serving as the vehicle for IBO delivery, the aCSF is (a) delivered alone to the brains of SHAM-operated controls; (b) the general tubing fluid between the IBO or aCSF/SHAM bolus and the injection syringe; and (c) a rinsing solution for the injection cannula/tubing/syringe system after cleaning and storage in alcohol. A 1 l volume of aCSF is made by the following three-part recipe: (1) Into flask A, place 500 ml of sterile distilled H2 O and add the following: NaCl KCl CaCl2 •2H2 O MgCl2 •6H2 O

8.66 g 0.224 g 0.206 g 0.163 g

(2) Into flask B, place 500 ml of sterile distilled H2 O and add the following: 0.214 g or Na2 HPO4 •H2 O (dibasic) Na2 HPO4 •7H2 O 0.358 g NaH2 PO4 •H2 O (monobasic) 0.027 g (3) Add flask A to flask B. This gives a 1 l solution with 150 μM Na, 3.0 μM K, 1.4 μM Ca, 0.8 μM Mg, 1.0 μM P, and 155 μM Cl. This solution has pH 7.4. Artificial CSF may be kept in a regular refrigerator in a covered flask. The somewhat more challenging work of preparing IBO in the aCSF vehicle is described in the following four steps, which require pH litmus paper, micropipettes, and Eppendorf tubes: (1) In the small vial in which the 5 mg IBO arrives, add 500 μl of Millipore sterilized aCSF. Use a button spinner to briefly agitate the solution in the container at this point and after subsequent additions of solution described next. (2) After adding the aCSF to the original container, the IBO does not completely dissolve. Now add 1 μl of 10 N NaOH (MW 40 g/mol) to help ionize the acid and facilitate


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solubility. Check the bottom of the container to see how much IBO remains undissolved; repeat these 1 μl additions of 10 N NaOH until only a few tiny IBO clumps are visible. (3) Now the IBO is nearly, but not totally, in solution, and the pH is still not quite physiologic (e.g., it is 12 rats) will lead to higher frequencies of PD-7 rats below the 15 g weight minimum. To address this possibility, and to anticipate the number of subjects available for surgery, it is recommended that litters be assessed at PD-3 to 5 and gendertyped and culled as needed (more culling will speed growth per animal). Gender-typing pups before PD-3 can be tricky but is quite accurate thereafter with a brief visual exam of pup’s genitalia (Fig. 1.4). Generally, compared to females, males will have a greater distance between anus and phallic prominence, and, no huge surprise, the males have a larger phallic prominence. On the day of surgery, it is important to balance randomized NVHL vs. SHAM assignments across and within available litters. Across litters, this is important for minimizing the effects of genetic heterogeneities between moms. Allowing for expected rates of attrition due to incorrectly placed lesions, this means generating NVHL vs. SHAM ratios (e.g., 5:3 NVHL/SHAM), as consistently as possible across litters. Within litters, even when

female

male

Fig. 1.4. Sketch of gender differences in genitalia in PD-3–7 pups.


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keeping within the 15–20 g weight range, it is important to optimize balance of lesion status by weight (e.g., ensure all the SHAM animals do not tend smaller). 3.2. Performing NVHL and SHAM Control Surgeries 3.2.1. Surgical Setup

On the day of surgery, setting up should include the following: Hypothermic anesthesia bucket: Prepare the anesthesia bucket as a rubber ice bucket filled with small ice chips/balls. The researcher’s fist is used to punch a central crater in the ice field 5–10 cm deep for pup placement. Cannula preparation: Tubing and syringes must be clean and patent with easy gliding motion of the syringe plunger. Between surgical sessions, the lines should be flushed with a tuberculin syringe loaded, first with 90% EtOH (4–5 ml squirted vigorously through a 25-gauge needle into the plunger-less Hamilton syringe end of the cannula) and second with aCSF flush (4–5 ml). During these flushes, one wants to see a thin but vigorous stream of fluid emerging from the injector tip. If the flow is slow, repeated flushing may be necessary, possibly with the use of cleaning wires provided with Hamilton syringes for removal of matter from inside the needles. The functioning cannula is then filled completely with aCSF free of air bubbles in the lines (although air bubbles may initially be used to check for good flow in the cannula as described above). As with the cleaning, this insertion of the “hydraulic” aCSF is accomplished by a more gentle injection of aCSF through the plunger-less Hamilton syringe with a tuberculin syringe. Now, for loading the injection bolus (e.g., IBO, aCSF, or dye), one first pulls back on the Hamilton’s plunger to create an air bubble of 0.2–0.3 μl volume as a mobile boundary between the aCSF hydraulic and the injection solution. This bubble not only prevents dilution of the IBO or dye into the hydraulic aCSF but also provides a visual marker to confirm that flow is occurring within surgeries, and for knowing for how much injection solution is left in the cannula over the course of multiple surgeries. Now with the indicator bubble in place, simply suck up IBO (from an Eppendorf tube), aCSF, or dye as needed. For a run of eight or more NVHL surgeries, we recommend filling the line with >7 μl by gentle backward plunging on the Hamilton syringe. If not anticipating a need for IBO for the rest of the session, return it to –80 freezer; if reloading later in the day, it may be stored on ice and under cover for 2–4 h in the surgical room. Now, mounting the cannula needle onto the stereotax, it is key to have the needle secured to the injector arm as close to the tip of the needle as possible for stability (Fig. 1.5), because the needle itself is somewhat unstably flexible. Also, the bevel of the injection needle should be squarely positioned facing the caudal aspect of the rat. After mounting the Hamilton syringe into the infusion pump, go ahead and manually waste some injection fluid out the injector needle tip

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no

yes

Fig. 1.5. Proper mounting of the injection needle on the stereotaxic arm.

to again confirm patency and ensure that the infusion pump and the syringe are correctly mechanically interfaced. Finally with no pup on stage, make one or two 0.3 μl infusions to verify proper function and volume delivery, and to generate a consistent balancing of hydraulic tension in the system. Wipe away the small infusate bead on the tip of the needle. Pre-op ready chamber: Properly selected pups (about six at a time, at the start of a run) should be on hand in a rat-housing tub. Each rat should be numbered with a marker on their back, with weight references, semi-random lesion assignments, and possibly gender ID noted on paper. Logistically, it is best to perform a series of rats (six or more) as IBO lesions and another series as SHAMS, to prevent from having to change cannula between surgeries. If two setups are used, they may be segregated for NVHL vs. SHAM surgeries. Post-op chamber: This may be a rat-housing tub, containing soft padding overlying a warming pad. 3.2.2. Inducing Hypothermic Anesthesia

Although unfamiliar to some investigators and animal research review committees, generalized hypothermia is safer, easier, and less expensive than chemical anesthesia for infant rats. The pup is placed in the ice crater for 15–20 min, with the cover placed over the ice bucket. During this induction, the pup should be checked every 5 min or so, as initially it will often succeed in crawling out of the crater to the edge where hypothermia occurs unevenly. In the 15–20 min window, the rat should be checked for complete immobilization and lack or respiration. This is achieved as early as 17 min. This induction should be rigorously timed. Leaving pups on ice for longer than 20 min significantly increases risk of death; taking them off too soon increases the risk of premature return of mobility during surgery, which is a major cause of lesion inaccuracies or unintended mechanical lesions in SHAM animals.


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Usually, 17–18 min on ice will provide 12–16 min of immobility, which in experienced hands is a sufficient time to perform the surgery. 3.2.3. Lesioning

The two major issues with the neonatal lesion surgery, as opposed to an adult stereotaxic surgery, are (1) the targets are much smaller and (2) the skull bone is far thinner and highly flexible. The former increases the challenge, while the latter provides both up and downsides. On the upside, there is no need for a bone drilling step; the injection needle tip itself is the drill. On the downside, the pup’s braincase is fragile and flexible, rendering head immobilization virtually impossible with traditional means. Thus, your target is very small, and to some extent, it will be moving, because anything (needles and fingers) that touches the pup will displace it after the stereotaxic coordinates are set. This has two important implications for the attitude of the researcher. First, compared to adult stereotaxic surgery, it has to be accepted that neonatal lesioning is much more of an acquired art or a skill (like bowling, dart throwing, and golf ). Second, like these sports, it takes practice to become reasonably proficient. Next we provide detailed guidelines on how the surgery should be approached, while Section 4.2 outlines approaches in practice lesioning. After induction of anesthesia, lesioning consists of three basic steps: (a) mounting the rat; (b) puncture and infusion; and (c) post-op recovery. Including anesthesia, all of these steps performed for a single rat take 40–55 min. However, when performing serial surgeries in assembly line fashion, one rat is undergoing anesthesia while another is being operated on, while another is in recovery. This shortens the procedure to 15–20 min per rat, or shorter with more than one setup. The remainder of this section assumes only one setup and is performed identically with IBO infusions (NVHL) or aCSF (SHAMs). (1) Mounting the rat: This is a crucial step in the process as mistakes here amplify the chance of inaccurate targeting. After the rat has been adequately anesthetized, place it squarely on its abdomen on the surgical platform with the aid of guide marks drawn on the platform. The rat’s tail should be pulled out straight behind and its four limbs should be spread out to the sides so that they do not sit under the rat’s neck and torso (Fig. 1.6). All this is to ensure that the body and the head sit on the platform in the most stable, central and square position (with respect to the stereotax arm movement). Now, using a scalpel or a sterile razor blade, a single incision is made down the middle of the rat’s head (1.5–2 cm long) so that bregma and the sagittal and coronal cranial sutures are observed. Be careful not to lacerate the veins or the tissue on the exposed skull. Spread out the

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Coronal Suture

Lamboidal Suture

Bregma

Saggital Suture

Fig. 1.6. Proper positioning and taping of pups on the stereotaxic stage.

width of the wound with your fingers to ensure adequate lateral room for insertion of the cannula needle into the skull. Now, using strips of freezer tape, secure the rat’s neck and head to the platform crosswise (Fig. 1.6). The caudal strip is placed first to stabilize the neck. Then, one hand is used to manually re-position the head so that it is flat on the platform and square with the stage (e.g., the sagittal suture is made parallel with the long-axis motion of the stereotaxic arm and non-rotated with respect to the vertical), while the other hand places the rostral strip of tape. Note that either or both the caudal and rostral pieces of tape can be placed just a bit over the open wounds to aid in keeping the wound open. Further, quick adjustments should now be made as needed to the tape and/or the rat’s positioning as necessary to maximize alignment of cranial landmarks with the stereotax (you will notice that the rat’s skull slides around very easily within the skin around the rat’s head). In making all these adjustments, it is important to view the animal from directly above, as a viewing angle off the vertical will result in a misaligned pup. (2) Puncture and infusion: Now using the stereotax, position the needle directly over bregma. In these infant rats, bregma may be quite difficult to see. Confirm that you have bregma


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by very gently taking the needle down to the skull surface. Carefully applying some pressure with the needle (do not puncture the sagittal vein!) will depress the soft tissue between the cranial plates giving contour to bregma and the cranial sutures. Looking at the skull from a sideways view so that the light is reflected from the skull can help reveal these contours. Once bregma is confirmed, write down the anterior–posterior (AP) and medial–lateral (ML) coordinates as given by the stereotax. Remember, ventral–dorsal (VD) coordinates are not needed because of the standardized depth needle band. The coordinates relative to bregma for the NVHL surgery are AP –3.0 mm and ML ± 3.5 mm (VD –5.0 mm). Move the needle using the stereotactic mechanism posteriorly 3.0 mm from bregma. Observe that as you move it back, it traces the sagittal suture. If it does not appear to follow it in this fashion, then you have got an alignment problem that needs to be fixed as described under the previous step (mounting the rat). It does not matter which side of the rat’s head you lesion first as long as you are consistent. Using the stereotactic mechanism and coordinates, move the needle 3.5 mm from the midline toward the side you choose to do first. Now you are ready to penetrate the skull. First use the needle as a punch to make a hole in the targeted spot on the skull. Holding the rat’s head steady with one hand (gentle pressure with thumb and index finger on either side), make a quick and shallow jab with the needle using the stereotax with the other hand. Now, keeping one hand in support of the rat’s head, slowly re-advance the needle into the hole until the standardized depth needle band touches (and somewhat depresses) the rat’s skull. The most likely complication with needle insertion (and a major source of inaccuracy) is rotation of the rat’s head to one side; gentle head support with the fingers and a properly targeted needle should keep this from happening. Once the needle is in, you should notice that the pressure of the needle band has forced blood out of the venous sinuses of the sagittal and lambdoid sutures. Slowly raise the needle just until the blood refills these veins. Now that the needle is in place, the infusion pump may be started. It should pump in 0.3 μl of fluid over the course of 2 min and 15 s (135 s). Once the infusion is complete, let the needle rest in place in the rat’s head for about 3 min to prevent backflow. For this duration of needle placement, it is recommended that a stopwatch set to 5 min be used to time from the start of the infusion to when the needle should be withdrawn. After 5 min of total needle placement, we now lesion the other side of the brain. In our experience, this second infusion is

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more likely to be off target than the first because the effects of pup motion and/or subtle misalignment, if present, are cumulative. Again, gently support the head with thumb and index finger of one hand, as this prevents cranial rotation during withdrawal of the needle. Still holding the head in this way after the needle is out, use the other hand to manipulate the stereotax so that the needle moves over 7.0 mm to the other side of the head (e.g., 3.5 mm to the other side of the sagittal suture). With good head support, the needle is now positioned above a spot that is symmetric with the AP and ML positions of the puncture hole on the contra-lateral side of the head. If when viewing the animal from directly above you do not see this symmetry, you may have to manually adjust the targeting just a bit (i.e., deviate from the exact coordinates) based on your line of sight only. This maneuver can be tricky but is sometimes necessary if inadvertent pup motion has occurred in prior steps, and it is often accurate with the eye of an experienced surgeon. Now, just as for the procedure on the other side of the head, and while providing head support, repeat the steps: punch and needle placement, timed infusion, and 3-min waiting period while the needle rests in the rat’s head. At the end of the second 3-min waiting period, the needle is withdrawn and the surgery is nearly complete. If during the surgery the rat begins to show movement (usually begins with back legs), put more pieces of ice over its back to maintain hypothermia. It is important to maintain immobility as much as possible during all phases of needle placement; movement of the head can make an IBO lesion too large or cause the SHAM surgery to make a mechanical lesion. After withdrawing the needle, carefully pull the tape off the rat. Take advantage of the remaining anesthetized state for using a mouse ear punch as needed to mark animals (one or two punches on either ear as NVHL, SHAM or probable failed attempt). It is best to close the wound after putting holes in the ears to prevent re-opening of the wound. Close the wound with a thin line of veterinary wound closure glue. (3) Post-operative recovery: Pups are placed in the post-operative tub for warming and monitoring. If a post-operative pup has not yet begun to stir, it should within the next 5 min. Animals receiving IBO may be observed to be more mobile compared to SHAM animals within this time frame. Occasionally, an IBO pup will die 5–10 min after becoming mobile, possibly in relation to improper ventricular spread of IBO that may impair respiration after recovery from hypothermia. Animals that die after showing little to no movement most likely had lethal hypothermia (an outcome


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that can be 100% avoided with proper timing of anesthesia induction). Proactive attempts to resuscitate pups with diminished or lost respiration are ill advised, since it may indicate a bad lesion, or extra-lesion damage (e.g., cerebral hypoxia). If after 15 min, the pups are respiring normally, motorically responsive to touch, have normal pink coloration, and dry wound closure cement (if used), they are ready for return to their litters. Ordinarily, rats are returned to their original litters. However if culling or other sources of attrition have greatly reduced litter sizes, adoption to other litters is reasonable, as long as litters remain balanced and equal numbers of NVHLs and SHAMs are adopted. 3.3. Lesion Confirmation

Lesion confirmation is a necessary step for all rats in a given experiment (e.g., including both NVHL and SHAM-operated animals). Bilateral success rates for NVHLs may vary (even for the same surgeon) from 50 to 100% between cohorts, and occasionally a SHAM will have enough needle track damage to warrant exclusion. Rats may be sacrificed at any age for this purpose, but consistent with most experimental designs, we describe methods general to adult rats (>PD-56).

3.3.1. Histology

After sacrifice (by deep anesthesia and guillotine) and brain removal, the brain is placed whole into an isopentane-filled cylinder (kept between –30 and –40◦ C on dry ice), for 20–40 s. The rock-hard brain is then removed with forceps and wrapped in an appropriately sized (∼12 cm × 12 cm) and labeled square of aluminum foil. The brains may then be stored in a –80◦ C freezer until cutting. For sectioning, aim to collect 6–8 coronal sections on a cryostat through most of the rostral–caudal extent of the hippocampus. Generally, sections for slides should begin when the dorsal– rostral blade of the hippocampus comes into view (∼–2.0 mm AP bregma (5) as shown in Fig. 1.7). Sections are collected for slides about every 400 μm. We have used 20-μm-thick sections, in which every 18th–20th section is put on slides, or 40-μm sections, with every 9th–10th kept. Four to six sections are mounted on each appropriately labeled slide. Before staining, mounted slides may be stored in a slide rack for several days in a –20◦ C freezer. For staining, allow slides to warm from cold storage at room temperature for about 5 min. Then, process slides through the following 17 solution steps, while allowing the dipping rack to drain off between emersions. Notably, only the steps marked by (∗ ) require an actual solution dish preparation; steps 4 and 6–9 may reuse solutions 1–3.

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Fig. 1.7. Appearance of dorsal blade of the hippocampus in coronal section; approximate area where first section for histological processing should be taken.

Dehydration (1) 70% ETOH (3 min)∗ (2) 95% ETOH (3 min)∗ (3) 100% ETOH (3 min)∗ (4) 100% ETOH (3 min) Fixing (5) 1:1 chloroform/100% ETOH (1 h to overnight) ∗ Rehydration (6) 100% ETOH (1–3 min) (7) 95% ETOH (1–3 min) (8) 95% ETOH (1–3 min) (9) 70% ETOH (1–3 min) (10) Calcite water (1–3 min)∗ (11) Calcite water (5 min)∗ Staining (12) Thionin solution (0.05%) (5–7 min)∗ (13) Calcite water∗ (14) 80% ETOH (1 min)∗ (15) 95% ETOH (1 min)∗ (16) 100% ETOH (1 min)∗ (17) Citrosolv (2 min)∗ (18) Coverslip with DPX (or equivalent)


3.3.2. Grading Lesions

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Investigators should examine brains from all animals in a given experiment, blind to the results of the completed study. It is recommended that brains be examined under both very low (4×–10×; magnifying glass) and low magnification (40×; microscope) to allow close visualization of the entire brain and a more detailed exam of the cellular architecture of the hippocampus. Both lesioned and SHAM rats should be examined. Many SHAMs will normally show trace evidence of needle track damage – and that is permissible, unless there is pronounced evidence of tissue distortion or destruction, suggesting a mechanical lesion. The coordinates for the NVHL target the area where the CA3 layers of the dorsal and ventral blades merge [in adult sections, at about −5.0 mm from bregma (5)] (Fig. 1.8). Lesioned animals are typically characterized by one or more of the following: (a) generalized enlargement of the lateral ventricle; (b) tissue atrophy and/or distortion of normally curvaceous cell layers in the ventral hippocampus (involving CA1, CA2, CA3, and subiculum) and/or the CA3 of the ventral extent of the dorsal hippocampus; (c) loss and disarray of individual neurons (e.g., decrease in thionin staining density in cell layers) within cell layers. Between subjects, it is to be expected (and is permissible) that lesion damage within the hippocampus is somewhat heterogeneous in size and exact location, much as histopathological studies of brains of schizophrenia subjects show within-group heterogeneity of morphological findings. However, the following parameters should also be used to disqualify rats and limit this heterogeneity. (1) Lesion size: Lesions can be too small or too large, and either extreme may alter the behavioral phenotype (6). Require evidence for IBO damage (not just needle tracks) in at least two serial sections. Conversely, the lesion should not be a total hippocampectomy or involve significant damage to the dorsal blade of the hippocampus as this structure has differential connectivity patterns and functionality compared to the ventral (more limbic) hippocampus (7). This outcome is easily ruled out by ensur-

Sham (normal)

IBO

Fig. 1.8. Primary target areas (cross hairs) of coordinates for NVHLs (left) and typical zone of tissue loss with a “perfect” bilateral hit in an NVHL (right).

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ing lack of extension of the lesion into the dorsal blade on the first section in the series. (2) Lesion location: The damage should be to the hippocampus proper; significant damage evident to the overlying cortex, underlying thalamus, or ventrally posited amygdala should be inspected for, and warrants disqualification. The NVHL lesion does cause some degree of generalized atrophy to its efferent/afferent structures (e.g., prefrontal cortex, temporal/entorhinal cortex, amygdala, dorsal hippocampus, and likely elsewhere) and so if noted on exam, such trends are not grounds for disqualification unless there is evidence of significant cell loss and/or gliosis in these structures under 40×. (3) Bi-laterality: Small lesions on one side and large on the other are not infrequent and are permissible as long as the lesions on each side meet the above criteria. However, a lack of bilateral lesions is an exclusion, representing either a lack of enough infusion volume on one side or an extrahippocampal infusion that may be hard to find.

SHAM

NVHL

Fig. 1.9. Example of published demonstration of serial maps showing smallest and largest extent of lesion damage in a study (8) and exemplary micrographs of SHAM vs. NVHL, with permission from Springer.


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For purposes of publication, it is typical for researchers to show anatomical maps of largest vs. smallest lesion extents of rats with appropriate lesions (e.g., from among all those not disqualified from a given study) along with a representative micrograph of a SHAM and NVHL brain (e.g., Fig. 1.9).

4. Notes: Problems and Pitfalls 4.1. Making Ibotenic Acid

4.2. Practice Lesioning

This obviously crucial step, if not done carefully and correctly, can cost serious time and money since histological lesion results are typically not assayed until well after rats reach adulthood (PD-56) and experiments are completed. The addition of NaOH for putting IBO into solution is a delicate matter and must proceed in a gradual and deliberate titration with frequent pH checks. The danger is that the pH can suddenly shoot higher than 8.5 from a prior reading of 25%, “bad acid” is a likely culprit, and a new batch should be mixed. To obtain a reasonable level of skill and accuracy, it is recommended that surgeons inexperienced with NVHLs initially practice lesions on 50–100 pups. Learning curves are more rapid if this is done in batches of 1–10 rats, with meticulous intra-surgery notes taken on each rat and batch sacrifices for lesion verification conducted within 24 h, if not immediately after a surgery. Keep a record of improving bilateral success rates vs. unilateral hits and total misses. Two major alterations to the general methods are important for practice lesioning: first, use Chicago Sky Blue (50 mg in 20 ml of aCSF) (or equivalent stain) as the infusate. Second, the histological processing involves sacrifice of rat pups [e.g., with lethal (>25 min) hypothermia, brain removal, isopentane-rapid freezing, and cutting on a cryostat only]. The lesion verification is done by visual inspection (with magnifying glass) of dye delivery while the brain is being sectioned on the cryostat – there is no need for slides or histology. Of note, the PD-7 rat brain and hippocampus do differ architecturally somewhat from the adult. After histology, the adult hippocampus appears to have two independent lobes: one dorsal that comes into view first (coronal sections cut caudally) and then a ventral that appears independently, then merges with the dorsal. In the pup, these parts, appearing darker than overlying cortex, seem more contiguous regardless of which section you are looking at. A good bilateral hit vs. common patterns of missed hits are shown in Fig. 1.10. In general, the key will be achieving consistent bilateral

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Bilateral Success

No

No

No

Fig. 1.10. Patterns of success and typical failures in practice lesioning using dye.

symmetry, alignment, and infusion depth. Create small cartoons for each result, compare with written notes, and adjust your technique accordingly. 4.3. SHAM Controls

It is debatable whether SHAM lesions are the best control for all studies. Mechanical damage or other complications of introducing a needle into the brain are likely to have at least some subtle long-term effect consistent with a mild lesion, potentially decreasing effect size in some NVHL vs. SHAM lesion comparisons. Although early piloting studies and some published results have shown little effect of SHAM–NVHL lesions compared to unoperated controls in a limited set of phenotypic measures, SHAM lesions of the neonatal medial prefrontal cortex do show trends approaching significance across several phenotypic measures compared to unoperated controls (9). Therefore, depending on the experiment and experimental justification, one or more alternative controls may also be used (e.g., unoperated pups not removed from litters; pups undergoing hypothermic anesthesia; and superficial wound opening only).

4.4. Cautions About Alternative Approaches

In developing or learning the methods for producing NVHLs, it may be tempting to try alternative modifications to the procedures. Here, we mention a few of these that we would advise caution in pursuing:


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Anesthesia: Although alternative forms of anesthesia (e.g., pharmacological) other than hypothermia have been used for NVHL surgery, we generally advise against these approaches. In our experience, hypothermia, when properly done, is safer, more predictable, cheaper, and cleaner. Using it arguably maintains the model as a more purely non-pharmacological model of schizophrenia, while other anesthetics if used this early in the developmental stage of the animal may have longterm effects and have themselves been used to produce models of schizophrenia (e.g., ketamine). Head stabilization: It may be tempting to create an apparatus for allowing greater head stabilization as an attempt to reduce the amount of practice and skill needed to make accurate lesions. In our experience, such attempts have amounted to wild-goose chases. While some groups appear to have successfully made form-fitting molds for stabilizing the rat’s body and head in the operative platform, we recommend against the use of metal probes, clamps, or ear bar-like instruments. The pup’s skull is simply too soft, flexible, and rotatable under the skin. Wound closure: The wound may of course be closed with wound closure glue, clips, or sutures. However, the latter unnecessarily adds time to the surgery and is susceptible to wound re-opening after return to litters. Mothers may also meddle with sutures and especially clips. Placing clips involves additional risk of injuring the pup’s skull, and they need to be removed a week or two later which unnecessarily distresses the animals. The only caution with glue is to make sure it is dry before returning pups to their litters.

Acknowledgments Compilation of this chapter was supported by NIDA K08 DA019850 (R.A.C.). The authors wish to thank Alena Sentir for her effort in rendering the original drawings into digital form. References 1. Lipska BK, Jaskiw GE, Weinberger DR. (1993) Post-pubertal emergence of hyperresponsiveness to stress and to amphetamine after neonatal excitotoxic hippocampal damage: a potential animal model of schizophrenia. Neuropsychopharmacology 9: 67–75.

2. Tseng KY, Chambers RA, Lipska BK. (2009) The neonatal ventral hippocampal lesion as a heuristic neurodevelopmental animal model of schizophrenia. Behav Brain Res 204: 295–305. 3. O’Donnell P, Lewis BL, Weinberger DR, Lipska B. (2002) Neonatal hippocampal

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damage alters electrophysiological properties of prefrontal cortical neurons in adult rats. Cereb Cortex 12:975–982. 4. Chambers RA. (2007) Animal modeling and neurocircuitry of dual diagnosis. J Dual Diagn 3:19–29. 5. Swanson LW. (2004) Brain maps: structure of the rat brain. 3rd ed. New York, NY: Elsevier. 6. Swerdlow NR, Halim N, Hanlon FM, Platten A, Auerbach PP. (2001) Lesion size and amphetamine hyperlocomotion after neonatal ventral hippocampal lesions: more is less. Brain Res Bull 55: 71–77.

7. Moser MB, Moser EI. (1998) Functional differentiation in the hippocampus. Hippocampus 8:608–619. 8. Chambers RA, Jones RM, Brown S, Taylor JR. (2005) Natural reward related learning in rats with neonatal ventral hippocampal lesions and prior cocaine exposure. Psychopharmacology 179: 470–478. 9. Schneider M, Koch M. (2005) Behavioral and morphological alterations following neonatal excitotoxic lesions of the medial prefrontal cortex in rats. Exp Neurol 195:185–198.

Chapter 1 - Springer Link

Chapter 1 - Springer Link

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