Approach to perfusion fixation
A major difference among studies that emerged was whether the investigators performed the perfusion fixation while the brain was still in the skull (i.e., in situ perfusion) or whether they removed the brain from the skull prior to performing the perfusion fixation (i.e., ex situ perfusion). There were two major subcategories for each approach. For the in situ approach, vessels were accessed either after making surgical incisions in the neck (or thorax) or after separating the head. For the ex situ approach, vessels were accessed either in the whole brain or in one isolated brain hemisphere. Two studies reported on multiple approaches. Istomin [
43] reported methods for both ex situ whole brain and in situ neck dissection approaches, whereas Waldvogel et al. [
96] reported methods for both ex situ whole brain and ex situ one-hemisphere approaches.
For the in situ approaches, one of the challenges described was difficulty perfusing the brain in the context of brain circulation deficits and/or brain trauma. Kalimo et al. [
45] reported that in two of the five brains that they attempted to fix via perfusion, there was no fixation noted when the brain was removed; in both of these cases, there was evidence to suggest premortem deficits in circulation to the brain. Böhm [
12], who performed the procedure on cadavers that had suffered injury to the head and brain, reported that the increased intracranial pressure resulting from brain death prevented cerebral perfusion throughout the internal carotid distribution. This was indicated by postmortem angiography that stopped at the intracranial internal carotid artery, which they called the “no-reflow phenomenon.” To mitigate this problem, Böhm [
12] opened the skull and capped the upper half of the brain prior to perfusion fixation. This problem appears to be mitigated by using the ex situ approach. For example, Sharma et al. [
79], who used the ex situ approach, reported perfusion fixation on brains donated from 5 individuals who had raised intracranial tension, or “pump brain,” prior to death. They found adequate or high-quality histology results when they did perfusion fixation on these brain samples.
Another challenge with the in situ approach is that it is more difficult to monitor perfusion fixation. Because the brain should harden during fixation, in an ex situ approach, it is possible to directly monitor fixation by applying pressure to the brain and noting resistance. In the in situ approach, the best monitoring method is likely fixation of the eyeball, which Donckaster et al. [
24] and Latini et al. [
53] both reported to be a suitable proxy for intracranial fixation. However, fixation of the eye may not always be completely reliable, due in part to the anastomosis between the external carotid and internal carotid through the ophthalmic artery. Kalimo et al. [
45] reported that even after clamping the external carotid artery, partial fixation of tissues in the external carotid distribution would occur unless digital pressure was applied to the inner supraciliary skin and perfusion fixation was kept to a short period of time. Finally, a practical downside of the in situ approach is that it can interfere with funeral and embalming practices. For example, Istomin [
43] noted that it was necessary to prepare the face of the cadaver prior to beginning the perfusion fixation, such as closing the eyes.
The in situ separated head approach was reported by 3 studies, all of which had the primary goal of surgical training. One consideration for the in situ separated head approach is the spinal level at which the head separation should be performed. Benet et al. [
9] performed the separation at vertebral levels C5-C7 to allow for sufficient exposure of the cervical vessels, while retaining the cervical spinal cord.
For the ex situ approaches, one of the challenges described is the mechanical damage and deformation that occurs while the organ is removed from its regular location in the skull. In the animal literature, mechanical postmortem trauma has been found to result in histological artifacts such as dark neurons [
44]. Investigators described several different approaches to minimize trauma. One approach is to suspend the brain in cloth; for example, Istomin [
43] reported using a hammock of dense fabric for holding the brain in place. Another approach is to bathe the brain in liquid; for example, Beach et al. [
7] placed the brain in phosphate-buffered saline. Beach et al. [
7] reported that of these two methods, the liquid bath solution may lead to less mechanical damage. Another challenge with the ex situ approach is that the arteries can be easily damaged while handling the brain, which will make subsequent perfusion more challenging or impossible. Beach et al. [
7] reported that when they removed the brain, they severed the carotid arteries so that there would still be long segments attached to the circle of Willis.
Regarding the ex situ one hemisphere approach, there are some special considerations. The process of cutting the brain introduces additional mechanical trauma that causes damage to the unfixed brain tissue and severs the arteries that supply the contralateral hemisphere, requiring additional artery ligations to prevent leakage of washout and fixative solution. Furthermore, the absence of collateral circulation from the contralateral circulation is likely to lead to worse overall fixation quality compared to the whole brain approach. In the process of cutting one hemisphere, it is also necessary to cut off the brainstem and cerebellum, with the result that these brain regions will not be perfusion-fixed because they are detached from the rest of the brain where the fixative is being perfused [
95]. As a result of these problems, the ex situ one hemisphere approach is typically performed only in cases where the other hemisphere needs to remain unfixed, to preserve the tissue for biomolecular or biochemical studies.
Taken together, there were four major approaches to brain perfusion fixation reported, each of which have reported benefits and downsides, although there is very little data on comparisons among them.
Brain donor exclusion criteria for perfusion
Many of the studies listed criteria for the inclusion of brain tissue in their studies; however, it was almost always unclear whether these exclusion criteria were specific to the perfusion fixation preservation procedure rather than overall inclusion in the study. The one exception is Adickes et al. (1996) [
2], in which cerebral vessel thrombosis or large intracerebral hemorrhages were both exclusion criteria specifically for perfusion fixation. In these cases, the investigators used immersion fixation. These exclusion criteria make biological sense, as these conditions are likely to interfere with flow through the cerebrovascular tree and therefore prevent adequate fixation.
While we did not identify any study that specifically noted that an extended postmortem interval (PMI) was an exclusion criterion for perfusion fixation, many of the studies reported the PMI range of the brain tissue used in their studies. The PMI range tolerated appeared to be associated with the goals of the investigators. On one extreme, Latini et al. [
53], who studied gross anatomy of the white matter, reported that they tolerated a PMI of up to 7 days, which was the longest PMI range we identified among the included studies. At the other extreme, Kalimo et al. [
45], who studied ultrastructure of brain parenchyma, used an “immediate autopsy” method such that their perfusion fixation procedure began within two minutes of death and the entire procedure was done within approximately 20 to 30 min after death. Another study of ultrastructure, by Suzuki et al. [
86], also required brain donors with a relatively short PMI of less than 5 h. They noted that autopsy cases after 5 h demonstrated worse preservation of the cytoplasm or cellular organelles, including vacuolar and liquefaction changes, which they attributed to autolysis. Somewhere in the middle of these extremes fell the majority of the light microscopy-based immunohistochemistry studies. For example, Beach et al. [
7] reported that they achieved “satisfactory” staining with PMIs of up to 18 h, although they noted that their immunohistochemistry results were best with brain tissue less than 12 h postmortem. As another immunohistochemistry example, Halliday et al. [
35] performed perfusion fixation on brains with PMIs of up to 35 h.
In summary, cerebral vessel thrombosis or large intracerebral hemorrhages were the only exclusion criteria specific to perfusion fixation. Several studies also suggested that a short PMI was preferred, with the PMI range tolerated depending on the type of the downstream study.
Vessels accessed for perfusion
Among the studies that we evaluated, there were many different choices in the vessels that they accessed for subsequent perfusion steps, which depended on the overall approach that they employed (Table
2). A key trade-off is ease of vascular access and technical perfusion quality versus the degree of dependence on intact collateral circulation for reaching more distant brain regions.
Table 2
Vascular access strategies reported by the included studies
| Ex situ, both hemispheres | Unilateral vertebral artery, bilateral carotid arteries | 18G cannula | Contralateral vertebral artery |
| Ex situ, one hemisphere | Internal carotid artery; if the PCoA was too small or not present, second cannula placed in the posterior cerebral artery | 18G plastic cannula | Basilar and contralateral cerebral arteries |
| In situ, head separated | Common carotid arteries, vertebral arteries, internal jugular veins | One-way urinary catheter (largest possible) | NR |
| Ex situ, whole brain | Bilateral internal carotid arteries, bilateral vertebral arteries or basilar artery | Plastic IV cannula | NR |
| In situ, head separated | Common carotid arteries, vertebral arteries, jugular veins | NR | NR |
| In situ, thoracic dissection | Aortic arch | Wide balloon catheter | NR |
| Ex situ, whole brain | Carotid and vertebral arteries | NR | NR |
| Ex situ, one hemisphere | Internal carotid artery, posterior communicating artery* | 20G peripheral catheter* | Basilar artery* and contralateral hemisphere arteries |
| In situ, neck dissection | Bilateral carotids, with or without vertebral arteries | Irrigation cannula | External carotids |
| Unclear | Carotid artery | NR | NR |
| Ex situ, whole brain | Bilateral internal carotid arteries and vertebral arteries* | Olive C cannula* | NR |
| Ex situ, whole brain | Carotid and vertebral arteries | NR | NR |
| Ex situ, whole brain | Bilateral internal carotid arteries and vertebral arteries | NR | NR |
| Ex situ, whole brain | Both internal carotids, if both PCoAs were sufficient diameter; One carotid and the basilar artery otherwise | NR | Non-cannulated arteries were ligated |
| Ex situ, whole brain | Internal carotid arteries and basilar arteries | NR | NR |
| In situ, neck dissection | Bilateral carotid arteries | NR | NR |
| In situ, neck dissection | Initial segment of the right internal carotid artery | Glass cannula | Right external carotid, both left carotid arteries, and vertebral arteries |
| In situ, neck dissection | Left or right common carotid artery | NR | NR |
| In situ, unclear approach | Internal carotid artery | NR | NR |
| Ex situ, whole brain | Bilateral internal carotid arteries | NR | NR |
| Ex situ, whole brain | Bilateral carotid arteries | NR | NR |
| In situ, neck dissection | Bilateral common carotid arteries | Polyethylene cannula (1/4″ outside diameter) | Vertebral arteries and internal jugular veins (intermittently clamped) |
| Ex situ, whole brain | Bilateral internal carotid and vertebral arteries | NR | NR |
| In situ, unclear approach | Unilateral carotid artery | NR | NR |
| Ex situ, whole brain | Blood vessels at the base of the brain and floor of the third ventricle (non-vessel) | NR | NR |
| Ex situ, whole brain | Bilateral internal carotid and vertebral arteries | NR | NR |
| Ex situ, whole brain | Bilateral internal carotid arteries and basilar artery | NR | NR |
| Ex situ, whole brain | Bilateral middle cerebral arteries | NR | NR |
| In situ, neck dissection | Left internal carotid artery | NR | NR |
| Ex situ, whole brain | Bilateral internal carotid arteries and the basilar artery | NR | After initial perfusion fixation, clamped vessels to isolate the hippocampus |
| In situ, head separated | Bilateral internal carotid arteries | One-way number 10 Foley urinary catheters | External carotid arteries |
von Keyserlingk 1984 [ 93] | In situ, neck dissection | Internal carotid artery, vertebral artery | NR | NR |
| Ex situ, whole brain | Basilar and internal carotid arteries | 21G winged infusion needles | Leaking vessels occluded |
| Ex situ, one hemisphere | Internal carotid, vertebral, and anterior cerebral arteries | 21G winged infusion needles | Leaking vessels occluded |
| Ex situ, whole brain | Internal carotid and vertebral arteries | Serum 1 needle* | NR |
All of the included studies attempted to perfuse the anterior circulation of the brain via the carotid artery distribution in some form; either via the common carotid artery or arteries, internal carotid artery or arteries, or the aortic arch. Waldvogel et al. [
96] also reported cannulation of the anterior cerebral artery in their ex situ one hemisphere approach. If only one side of the two carotid arteries is cannulated for perfusion, then interhemispheric collateral circulation will likely provide some fixative to the other hemisphere via the anterior communicating artery [
55]. However, the perfusion quality in that hemisphere will be limited, especially if the anterior communicating artery is absent or hypoplastic [
78]. In the in situ approach, if the internal carotid was cannulated, several of the investigators (Table
2) also clamped the external carotid to prevent shunting of perfusate to the often lower-pressure external carotid circulatory distribution, as opposed to the brain.
Slightly more than half (20/32 or 62.5%) of the included studies reported consistently cannulating vessels in the posterior circulation in some form; either the vertebral artery or arteries, basilar artery, posterior cerebral artery, or the aortic arch. The remainder of the studies either did not focus on brain regions supplied by the posterior circulation or relied on collateral circulation from the anterior to the posterior circulatory system. Collateral circulation via the posterior communication arteries is not intact in approximately one-fifth of people [
102], although some degree of leptomeningeal collateral circulation may still be present [
73]. Notably, the ability to visualize the posterior communicating arteries directly is an advantage of the ex situ approach, as the likely amount of collateral circulation through the circle of Willis can be visually assessed and the vessels to perfuse chosen accordingly (performed by Insausti et al. [
42] and Adickes et al. (1997) [
1]).
For obvious reasons, it is technically easier to cannulate fewer arteries, and this also decreases the time interval for tissue degradation prior to the initiation of washout and fixation. Cannulating more arteries also potentially affects perfusion quality within each one of the arteries when using a perfusion setup with a tube splitter to distribute the perfusate, as was used in Beach et al. [
7]. This is because perfusion flow will distribute to the lowest pressure arteries, and cannulating a low-pressure artery that distributes fixative to a less important region of the brain may lead to worse quality fixation in a more important region of the brain. Finally, one of the advantages of the ex situ approach is that it is easier to access more blood vessels on the ventral surface of the brain without requiring more extensive neck dissection to access the vertebral artery. Relatively more of the studies using the ex situ than the in situ neck dissection approach reported consistently cannulating at least one artery in the posterior circulatory system (Table
2).
One study, Sharma et al. [
79], reported perfusion fixation via the lateral ventricles using the ex situ approach, in addition to the blood vessels. This method likely allowed for improved fixation of periventricular brain structures such as the hypothalamus. The lateral ventricular perfusion method was also used with good reported results by Toga et al., who used an in situ approach and was not identified by our formal search methods [
89]. This study found that their intraventricular delivery system led to better and more uniform fixation preservation quality than perfusion of fixatives through the carotid and vertebral arteries. They speculated that this was due to erratic blood clot formation during the postmortem interval.
Torack et al. [
90] reported a unique procedure in an attempt to isolate the hippocampus as a target for perfusion fixation. They first perfused through the internal carotid arteries and the basilar artery. Next, they clamped the middle cerebral artery distal to the anterior choroidal artery and the posterior cerebral artery distal to the posterior choroidal arteries. Following these occlusions, the perfusion fixation should have been more targeted to the hippocampus.
The main goal of vascular access points in perfusion fixation is to perfuse a large portion of the brain with little damage to the tissue. The studies that were able to successfully cannulate the anterior circulation as well as the posterior circulation would likely perfuse the largest amount of brain tissue. We are unable to determine if the quality of the tissue isolated from brains with different perfusion access protocols is significantly different.
Washout solution used
Slightly more than half (20/35 or 57%) of the included studies reported using a washout solution prior to perfusion fixation (Table
3). This step aims to remove clots, blood cells, and other intravascular debris to improve flow of fixative, although it comes at the cost of increased procedural complexity and a longer delay prior to fixation. Adickes et al. (1997) [
1] did not use a “pre-perfusion” or washout step with saline because it would make the procedure more burdensome on staff. Donckaster et al. [
24] only used their washout solution in cases with a PMI of more than 12 h prior to the initiation of the procedure, with the goal of preventing the fixation of blood clots. Of the studies that employed a washout step, saline or phosphate-buffered saline were the most common base washout solutions used, while two of the studies used mannitol, and one study used Ringer solution.
Table 3
Washout solutions used by the included studies
| Warm tap water | NR | Syringe (60 ml) | NR | 2–4 l | NR | NR | Until water flow was clear (clot/debris removal) |
| Ice cold PBS | NR | Pump | 10–20 min | 1 l | 50–100 ml/min | NR | NR |
| Isotonic saline | NR | NR | NR | NR | NR | “Low pressure” | Until contralateral outflow was clear |
| Ringer solution in 0.2 M phosphate buffer (pH 7.5) | Rheomacrodex (Dextran 40) | Gravity | 5–10 min | 5 l | 500–1000 ml/min | NR | Until blood and blood clots were washed away |
| 0.15 M PBS (pH 7.2) | NR | Pump | NR | 1 l | NR | NR | NR |
| Mannitol | Warm heparin | Gravity | NR | 250 ml | NR | NR | NR |
| Physiological saline | NR | NR | NR | NR | NR | NR | NR |
| NaCl 0.9% | NR | Gravity* | NR | NR | NR | 147.4 mmHg (height of 2 m*) | NR |
| 20% Mannitol | Heparin | Gravity* | NR | 250 ml | NR | 147.4 mmHg | NR |
| 0.1 M Sodium phoshate (pH 7.4) | 1% sodium nitrite | Pump | NR | 5 l | NR | “Normal mean arterial pressure” | NR |
| PBS | NR | NR | 33 mins | 4 l | 120 ml/min | NR | NR |
| Saline at 4 °C | Heparin, 10,000 units | NR | 20 mins | 2 l | 100 ml/min | NR | NR |
| Saline | NR | Gravity or Syringe | NR | NR | NR | 150 mmHg | Clear fluid flow from the veins |
| NaCl 0.9% | NR | Gravity | <= 5 mins | NR | NR | NR | NR |
| 0.01 M PBS (pH 7.4) | NR | NR | NR | NR | NR | NR | NR |
| 0.01 M sodium-PBS (pH 7.4) | NR | Pump | NR | 1 l | NR | NR | NR |
| Ice cold PBS (pH 7.4) | NR | NR | NR | 2 l | NR | NR | NR |
| PBS | NR | NR | NR | 180 ml (60 ml in each vessel) | NR | NR | NR |
| Saline | NR | Gravity | NR | 3 l | NR | 110 mmHg (height of 1.5 m) | Until no visible blood or clots drained from the IJVs |
| PBS (pH 7.4) | 1% sodium nitrite | Pump | 15 mins | 0.5 l | ~ 33 ml/min | NR | 15 min or until the brain is cleared of blood |
| Physiological saline | 0.33% heparin | Gravity | 30 mins | 1.5 l | 50 ml/min | NR | NR |
Published perfusion fixation methods for laboratory animals often start while the animal is anesthetized [
30]. This protocol prevents substantial premortem and postmortem clot formation [
36], which means that the major purpose of the washout solution is to remove blood cells from the vessels. On the other hand, in postmortem human brain perfusion fixation, there is frequently an abundance of blood clots that limit perfusion quality [
22]. This means that in addition to washing out the cells, the washout step is often used by investigators to also decrease the clot burden by driving them out with pressure. Böhm [
12] noted that the washout step removed most clots that had formed postmortem, while clots that were formed premortem could only be washed out if a higher perfusion pressure was employed. Notably, the goal of Böhm [
12] was to
preserve premortem clots for forensic purposes, whereas studies using perfusion fixation to study brain parenchyma typically aimed to remove clots in order to improve perfusate flow and resulting fixation quality.
In addition to mechanically removing blood clots via perfusion pressure, another approach is to degrade or inhibit clots enzymatically. Four of the studies added the anticoagulant heparin to their washout solution, which may help to limit the spread of blood clots (Table
3). One of the studies, Böhm [
12], reported the occasional use of dextran 40, which also has antithrombotic properties [
74].
Two of the studies, Halliday et al. [
35] and Waldvogel et al. [
96], reported the addition of sodium nitrite to the washout solution. Sodium nitrite may help to dilate blood vessels and has been found to improve perfusion fixation quality in animals [
71].
The volume of the washout solution varied considerably, from as little as 180 ml to as much as 5 l. Several of the studies also reported performing the washout step until the venous outflow was clear of blood, clots, or debris.
One potential problem with the use of a washout solution in brain perfusion fixation is that it may induce brain edema. In animal studies it has been shown that perfusing too much saline into the brain (e.g., one liter) can cause edema [
11]. The edema induced may be related to the osmotic concentration of the washout solution. Consistent with this, Benet et al. [
9] found that washing out with an isotonic saline solution rather than tap water led to decreased tissue edema. Grinberg et al. [
34] compared a hyperosmolar solution of 20% mannitol with a solution of 0.9% NaCl, finding that 20% mannitol led to substantially less brain swelling. Böhm [
12] also used a hyperosmolar washout solution (680 mOsm) composed of Ringer solution in 0.2 M phosphate buffer.
Overall, the majority of articles included a washout step, most commonly using 1–5 l of saline as the base washout solution. The additives used and the precise procedure reported differed widely, and there were few comparisons between methods.
Fixative solution used
Consistent with its widespread use throughout pathology and histology, formaldehyde was a component of the fixative used in almost all studies. The only exceptions were one condition in Grinberg et al. [
34] that employed 70% ethanol only (which did not lead to successful fixation) and 3 studies that used glutaraldehyde only (Table
4). Some studies used paraformaldehyde, which is a polymerized storage form of formaldehyde, while others used formalin, which is a form of formaldehyde that includes methanol to inhibit polymerization. 10% formalin is composed of 3.7% formaldehyde with around 1% or less of methanol [
88]. Paraformaldehyde typically requires depolymerization via heating and/or sodium hydroxide prior to use, thus adding another setup step that adds complexity and will potentially prolong the interval prior to the initiation of the procedure [
47]. The addition of methanol in formalin keeps the formaldehyde depolymerized and avoids its precipitation.
Table 4
Fixative solutions reported by the included studies
| 10% buffered formalin | NR | Gravity | 15–20 min | 2 l | 100–133 ml/min | 75.6 mmHg (height of 1 m) |
| 10% buffered formalin | Phosphate | Gravity | 15–20 min | 2 l | 100–133 ml/min | 75.6 mmHg (height of 1 m) |
| Formaldehyde 37% and ethyl alcohol 10% | NR | Syringe (60 ml) | NR | NR | NR | NR |
| 4% paraformaldehyde (ice cold) | 0.1 M phosphate buffer (pH 7.4) | Pump | 40–80 min | 4 l | 50–100 ml/min | NR |
| 10% formaldehyde | NR | NR | NR | 0.7 l | NR | NR |
| Custom solution: ethanol 62.4%, glycerol 17%, phenol 10.2%, formaldehyde 2.3%, and water 8.1% | NR | NR | NR | 0.7 l | NR | NR |
| 2% glutaraldehyde | 0.2 M phosphate buffer | Gravity | 5–10 min | 5–10 l | ~ 1000 ml/min | 25.7–47.8 mmHg |
| 4% paraformaldehyde | 0.15 M PBS (pH 7.2) | NR | NR | 3 l | NR | “Normal mean arterial pressure” |
| 20% formalin | NR | Gravity | NR | 5 Ll | NR | NR |
| Cajal fixative: formalin and ammonium bromide | NR | NR | NR | 900 ml (300 ml in children < 12 years old) | NR | < 200 mmHg |
| 10% formalin | NR | NR | NR | NR | NR | NR |
| 2.5% formaldehyde, 6% isopropyl alcohol, 1% glycerin | NR | NR | NR | NR | NR | NR |
| 10% formalin | None | Gravity | NR | 5 l | NR | 147.4 mmHg (height of 2 m*) |
| 20% formalin | None | Gravity | NR | 5 l | NR | 147.4 mmHg |
| 70% ethanol | None | Gravity | NR | 5 l | NR | 147.4 mmHg |
| Acetic acid-alcohol-formalin | None | Gravity | NR | 5 l | NR | 147.4 mmHg |
| 4% formaldehyde, 2% picric acid; followed by 10% sucrose in fixative | 0.1 M sodium phosphate | Pump | NR | 10 l fixative only; 4 l 10% sucrose in fixative | NR | “Normal mean arterial pressure” |
| 4% paraformaldehyde | 0.1 M phosphate buffer | NR | 83 mins | 10 l | 120 ml/min | NR |
| 4% paraformaldehyde (4 °C) or 4% paraformaldehyde, 0.02% picric acid (4 °C) | NR | NR | 120 mins | 4 l or 8 l | 33 or 67 ml/min | NR |
| 10–12% formalin | Neutral buffered | Syringe or Gravity | NR | NR | NR | 150 mmHg |
| 1.0% paraformaldehyde, 2.0% glutaraldehyde (37 °C) | 0.1 M cacodylate (pH 7.4) | Gravity | NR | 1.5 l (adult), 0.7 l (newborn) | NR | 132 mmHg |
| 12% formalin | NR | Infusion device (compressed air mechanism)* | 15–20 min | 2 l | 100–133 ml/min | 1500 mmHg (200 kPa) |
| 4% paraformaldehyde, 0.2% picric acid, and 0.1% glutaraldehyde | 0.1 M phosphate buffer (pH 7.4) | NR | NR | NR | NR | NR |
| 4% formalin | 75 mM phosphate buffer (pH 7.0) | NR | NR | NR | NR | NR |
| 4% formalin, 1% glutaraldehyde | 0.1 M phosphate buffer (pH 7.4) | NR | NR | 400 ml | NR | 100 mmHg |
| 10% buffered formalin | NR | NR | NR | NR | NR | 100 mmHg |
| 4% paraformaldehyde, 0.1% glutaraldehyde | 0.1% phosphate buffer (pH 7.4) | NR | NR | NR | NR | NR |
| 10% formalin | Neutral buffered | Gravity | 60 mins | 12–14 l | 200–233 ml/min | 75.6 mmHg (height of 1 m) |
| 4% paraformaldehyde, 0.1% glutaraldehyde (ice cold) | 0.1 M phosphate buffer (pH 7.4) | Pump | 15 mins | 1 l | 70–80 ml/min | NR |
| Alcohol 80% 9 parts, formalin 4% 1 part | NR | NR | NR | NR | NR | NR |
| 20% formalin | Neutral buffered | NR | NR | NR | NR | NR |
| 2.5% glutaraldehyde containing 0.2 M sucrose | 0.1 M phosphate buffer (pH 7.4) | NR | NR | NR | NR | NR |
| 4% paraformaldehyde, 0.2% glutaraldehyde | PBS | NR | 90 mins | 6 l | 67 ml/min | NR |
| 2.5% glutaraldehyde | Phosphate buffer (pH 7.4) | NR | 5–10 min | NR | NR | NR |
| 2% glutaraldehyde, 1% paraformaldehyde (pH 7.2) | 0.1 M sodium cacodylate | NR | NR | 0.7 l | NR | NR |
| 4% paraformaldehyde (4 °C) | 0.1 M phosphate buffer (pH 7.4) | NR | 30 mins | 1.68 l (560 ml in each artery) | 50 ml/min | “40 lbs. of pressure” |
| 10% formaldehyde | NR | Gravity | 60 mins | NR | NR | 110.4 mmHg (height of 1.5 m) |
von Keyserlingk 1984 [ 93] | 1% paraformaldehyde, 1% glutaraldehyde, 1.65% potassium dichromate | 0.1 M cacodylate buffer (pH 7.4) | NR | NR | 5 l | NR | NR |
| 15% formalin | 0.1 M phosphate buffer (pH 7.4) | Pump | 30–45 min | 2 l | ~ 33 ml/min | NR |
| 4% paraformaldehyde, 0.05% glutaraldehyde, and 0.2% picric acid | 0.1 M phosphate buffer | Gravity* | 90–120 min | 4–5 l | 33–56 ml/min | NR |
Twelve of the studies employed glutaraldehyde in the perfusion solution, at various concentrations ranging from 0.05% in Welikovitch et al. [
99] to 2.5% in Shinkai et al. [
81] and Suzuki et al. [
86]. In general, adding glutaraldehyde to the fixative solution allows for improved tissue morphology preservation for electron microscopy [
67], at the cost of decreased immunogenicity of antigens for immunohistochemistry [
47]. However, at lower concentrations of glutaraldehyde, such as the 0.05% used in Welikovitch et al. [
99], its effects on antigenicity are likely to not be as pronounced, and it likely acts primarily to slightly improve tissue morphology.
In addition to formaldehyde and glutaraldehyde, some investigators have used other fixatives. Picric acid, also known as 2,4,6-trinitrophenol, was used by Halliday et al. [
35] (2%), Insausti et al. [
42] (0.02%), Lin et al. [
57] (0.2%), and Welikovitch et al. [
99] (0.2%). Picric acid has been found to improve preservation of immunogenicity compared to aldehyde fixation alone [
82], although safety concerns make this fixative less desirable due to its explosive properties.
Pakkenberg et al. [
70] used a solution made up of 9 parts 80% alcohol and 1 part 4% formalin, which fixed the tissue to a quality sufficient for counting the number of nucleoli in the cortex, but also led to 20% volume shrinkage. This is consistent with the dehydrating effect of alcohol fixatives [
39]. Other studies that used alcohol in their fixative solutions included Feekes et al. [
28], Grinberg et al. [
34], and Benet et al. [
9].
Two of the studies used sucrose as a component of their perfused fixative solution, Shinkai et al. [
81] and Halliday et al. [
35]. The addition of sucrose might help to optimize the osmotic concentration of the perfusate [
13,
98] and/or to act as a cryoprotectant to prevent tissue morphology changes due to ice damage during sectioning with a freezing microtome.
Donckaster et al. [
24] perfused Cajal fixative, which consists of formalin and ammonium bromide. The addition of ammonium bromide is thought to facilitate silver staining of neural cells [
52]. von Keyserlingk et al. [
93] perfused 1% paraformaldehyde, 1% glutaraldehyde, and 1.65% potassium dichromate. The addition of potassium dichromate has been found to aid in the fixation of lipids [
38], which is consistent with the focus of von Keyserlingk et al. [
93] on myelin ultrastructure.
Benet et al. [
9] used a custom fixative composed of ethanol 62.4%, glycerol 17%, phenol 10.2%, formaldehyde 2.3%, and water 8.1%, which they compared to a fixative with 10% formaldehyde for use in surgical simulation. They concluded that the custom fixative was superior for surgical simulation, in part because it caused less hardening and therefore allowed for more realistic tissue retraction.
Grinberg et al. [
34] compared four different fixatives in their study. They found that perfusion of 20% formalin and acetic acid-alcohol-formaldehyde both led to efficient fixation of deep brain structures, while 10% formalin did not, and 70% ethanol did not harden at all. However, they found that the acetic acid-alcohol-formaldehyde fixative led to dissolution of myelin, while 20% formalin did not.
The fixative vehicle or buffer can also have important effects on tissue preservation [
16]. The most common buffer in the studies we identified was phosphate buffer, which was reported in 19 of the studies. Phosphate buffer can be titrated to maintain an approximately neutral pH, at which point the fixative solution can also be called “neutral-buffered.” One of the most important aspects of the buffer is the molarity, which is thought to be the major driver of the osmotic concentration of the fixative solution [
14]. Although there is some controversy on this point, aldehyde fixatives themselves are generally not considered major drivers of the osmotic concentration, as they easily cross semipermeable cell membranes, and therefore do not exert a sustained osmotic force [
37]. As a result, the osmotic concentration of the fixative vehicle is called the effective osmotic concentration. Hypertonic fixative solutions can cause grossly shrunken brain tissue and cell shrinkage, whereas hypotonic solutions can cause edema and resistance to flow in the perfusion procedure [
77].
It would be convenient to be able to identify the optimal vehicle osmotic concentration that would minimize osmotic tissue changes. However, Böhm [
12] pointed out that the redistribution of fluids and ions during hypoxia makes it difficult to identify this optimal osmotic concentration in the postmortem state, which is consistent with more recent evidence [
46,
51]. To study this empirically, Böhm [
12] used fixative solutions with multiple different osmolarities, finding that a mildly hypertonic solution with a total osmotic concentration of 500 mOsm and an effective osmotic concentration of 300 mOsm led to the best fixation quality in their study.
Several of the included studies manipulated the temperature of their fixative solution prior to perfusion. Beach et al. [
7] cooled their fixative solution to be “ice-cold,” while Torack et al. [
90] and Insausti et al. [
42] cooled their fixative solution to 4 °C. Lower temperatures can help to inhibit metabolism and thereby mitigate tissue degradation, although it has also been reported to cause vasoconstriction [
29]. One study, Kalimo et al. [
45], perfused their fixative at the elevated temperature of 37 °C, which has been suggested to facilitate vasodilation and improve perfusion flow [
29].
Taken together, 1–10 l of phosphate-buffered formaldehyde was the most common fixative solution perfused. The most important determinants of the fixative are the assay of interest and the tissue or cell type of interest (e.g. neurons or myelin). The choice of fixative buffer is an important way to balance tissue shrinkage and swelling while the fixative is being perfused and can affect fixation quality.
Driving perfusate and perfusion pressure
The three major methods for driving the flow of solution during perfusion are syringes, gravity, and perfusion pumps. All three methods were reported by the included studies: 2 studies reported using a syringe, 8 studies reported using gravity, and 4 studies reported using a pump (Table
4). The majority of studies did not report their drive method. Upsides of a syringe are that it is easier to inject a specific amount of fluid in each vessel, while it is more difficult to control flow rate and pressure.
From the perspective of a perfusion circuit, the included studies were open-circuit in that they did not describe using a method for re-introducing the outflow of the perfusate back into the vessels. In the in situ approaches, the perfusate typically drained from the internal jugular veins after flowing through the carotid and/or vertebral circulatory systems. In the ex situ approaches, the perfusate would be expected to drain from the cerebral veins and/or ruptured vessels below the isolated brain, for example into a container.
A major trade-off in setting the perfusion pressure is that too high of a perfusion pressure may lead to a higher risk of vessel rupture [
76], while too low of perfusion pressure may lead to incomplete perfusion, decreased clot removal, and decreased tissue penetration of the fixative [
17]. In laboratory animals, investigators often suggest that perfusion pressure should be maintained at roughly the same pressure that it was during life, which is called physiologic pressure [
25,
30]. Consistent with this, Halliday et al. [
35] and Coveñas et al. [
20] reported that their perfusion pressures were “normal mean arterial pressure.” Böhm [
12] kept their perfusion pressure lower, in the range of 25.7 mmHg (35 cm H
2O) to 47.8 mmHg (65 cm H
2O), because they were concerned that the endothelium is less stable postmortem than it is while the person is alive. However, Latini et al. [
53] used the supraphysiologic pressure of 1500 mmHg (200 kPa) to study white matter anatomy, and they were able to preserve and dissect white matter blood vessels of submillimeter size.
Techniques using syringes, gravity, and perfusion pumps have all been employed to drive perfusion flow at a variety of different pressures. However, there were no studies that made comparisons between these alternative methods or identified an optimal perfusion pressure range for a particular application.
Postfixation procedures
In the context of perfusion fixation, “postfixation” refers to immersion fixation of the tissue sample for some amount of time following the initial perfusion, either in the original fixative or in a new fixative solution. The procedure for postfixation depends on whether the perfusion fixation was perfused in situ or ex situ (Table
5). If in situ, then the brain was often left in the skull for some amount of time to allow for fixative diffusion prior to removal. This time period ranged from 1 h in McKenzie et al. [
64], 1 to 2 h in Kalimo et al. [
45], and 2 h in von Keyserlingk et al. [
93], to 48 h in Latini et al. [
53].
Table 5
Postfixation procedures reported by the included studies
| NR | 10% buffered formalin | Phosphate | NR | 1 day (if postfixed)a |
| Cut into 1–1.5 cm-thick sections | 10% buffered formalin | Phosphate | NR | 5–6 h |
von Keyserlingk 1984 [ 93] | Brain left in skull for 2 h, then removed and dissected | 1% osmium tetroxide | 0.1 M sodium cacodylate | NR | 2 h |
| NR | 10–12% formalin | Neutral-buffered | NR | NR |
| NR | 4% paraformaldehyde | 0.1 M phosphate buffer (pH 7.4) | 4 °C | NR |
| NR | 1:10 dilution of 10% formaldehyde | NR | 5 °C | > = 2 days |
| NR | 1:10 dilution of 10% custom solution (ethanol 62.4%, glycerol 17%, phenol 10.2%, formaldehyde 2.3%, and water 8.1%) | NR | 5 °C | > = 2 days |
| Cut into 1 cm-thick coronal sections | Paraformaldehyde or formalin | 0.1 M phosphate buffer | NR | NR |
| NR | 4% paraformaldehyde | 0.15 M PBS (pH 7.2) | 4 °C | 30 days |
| NR | 20% formalin | NR | NR | 3 weeks |
| Brain removed | Cajal fixative: formalin and ammonium bromide | NR | NR | 4 days |
| NR | Same fixative as was used for fixation | NR | NR | NR |
| Dissection of brainstem | NR | NR | NR | <= 24 h |
| Dissected into slabs approximately 1 cm thick | 4% paraformaldehyde | NR | NR | 48–72 h |
Kalimo 1974 [ 45] (electron microscopy) | Brain left in the skull for 1 to 2 h after perfusion fixation, then removed, then samples dissected for EM | 1.0% paraformaldehyde, 2.0% glutaraldehyde | 0.1 M cacodylate (pH 7.4) | NR | Overnight |
Kalimo 1974 [ 45] (histology) | Same as above | 10% formaldehyde | NR | NR | 10 days |
| Brain extracted from the skull 48 h after perfusion | 10% formalin | NR | NR | 24 h |
| NR | 4% paraformaldehyde | 0.1 M phosphate buffer (pH 7.4) | 4 °C | Overnight |
| Brain removed from skull | 4% paraformaldehyde | 0.15 M Sørensens phosphate buffer (pH 7.4) | 4 °C | 2 weeks |
| NR | 4% formalin | 0.1 M phosphate buffer | NR | > = 3 days |
Masawa 1993 [ 59] (electron microscopy) | From postfixed tissue, tissue blocks were cut and buffer washed | 1% osmium tetroxide solution | NR | 4 °C | 90 min |
| NR | 4% paraformaldehyde | NR | NR | 2–3 days or until the pink color of unfixed erythrocytes was gone |
| Waited 1 h after perfusion fixation, then the skull was opened, and the brain was removed | Formalin | Neutral-buffered | 4 °C | NR |
| Brain removed from skull | Alcohol 80% 9 parts, formalin 4% 1 part | NR | NR | 3 weeks |
| Brain suspended in a bucket | 20% formalin | Neutral-buffered | NR | 1–4 days |
| Cut into 2 mm-thick tissue blocks | 2.5% glutaraldehyde containing 0.2 M sucrose | NR | NR | 4–8 h |
| Brain halved sagittally and sliced into 10 mm coronal blocks | 4% paraformaldehyde | PBS | 4 °C | 2 days |
| Dissected bifurcations of the first temporal branches of the middle cerebral arteries | 2.5% glutaraldehyde | NR | NR | 4 h |
Tanaka 1975 [ 87] (electron microscopy) | Samples taken from various regions of the brain | 1.0% osmium tetroxide | NR | NR | NR |
Tanaka 1975 [ 87] (histology) | Rest of the brain | 8.0% formaldehyde | NR | NR | NR |
| Hippocampus and entorhinal cortex was isolated and sectioned into 0.5 cm thick slices | 4% paraformaldehyde +/− 1% Bouin’s solution (picric acid, acetic acid, and formaldehyde) | NR | NR | 48 h |
| Brain removed from skull | 10% formaldehyde | NR | NR | 2 weeks |
| NR | 15% formalin | 0.1 M phosphate buffer (pH 7.4) | NR | 6–12 h |
| Dissected out the medial temporal lobe | 4% paraformaldehyde and 0.2% picric acid | 0.1 M phosphate buffer | NR | Overnight |
Many of the studies reported cutting the brain prior to additional postfixation; for example, in Nakamura et al. [
69], the tissue was cut into 1–2 cm-thick coronal blocks. Perfusion-fixed tissue is harder and therefore easier to cut than fresh tissue. Cutting the tissue makes the subsequent immersion fixation process faster because there is a shorter distance for the fixatives to diffuse, with the obvious issue of damaging tissue at the cut interfaces.
There was a wide range of time frames used for postfixation, ranging from 4 h in Suzuki et al. [
86] and 5–6 h in Adickes et al. (1997) [
1] to 3 weeks in de Oliveira et al. [
23] and Pakkenberg et al. [
70] and 30 days in Coveñas et al. [
20]. How long investigators chose to postfix for may depend in part on their perception of the quality of their perfusion fixation. One major advantage of postfixation is that it will allow for fixation even in regions of the brain where perfusion has been minimal or absent, for example as a result of persistent blood clots.
A key trade-off in the length of postfixation is that longer amounts of time will lead to better fixative penetration of deeper regions of the brain or tissue block, while it may also lead to over-fixation and decreased antigenicity in the outer regions of the brain (i.e., the cerebral cortex) or tissue block. As a result, a significant disadvantage of a long period of postfixation is that immunohistochemical staining and quantification will result in variable gradients across the tissue section. However, these gradients can be minimized by pre-processing steps that cut the tissue into smaller sections prior to postfixation. For example, Shinkai et al. [
81] cut the tissue into 2 mm sections and Torack et al. [
90] cut the tissue into 5 mm sections prior to postfixation.
The majority of the studies used the same fixative for perfusion fixation and postfixation. One exception is glutaraldehyde fixation studies, which typically omitted it from the postfixative, likely in order to mitigate further antigen masking. Another exception is three studies that prepared tissue samples for electron microscopy, Tanaka et al. [
87], Masawa et al. (1993) [
59], and von Keyserlingk et al. [
93], which postfixed in osmium tetroxide, a fixative that stabilizes the ultrastructure of lipids and cell membranes [
26].
In summary, postfixation is used commonly and it allows investigators to compensate for the possibility of poor perfusion quality. There was a wide range of postfixation procedures reported, ranging in time from a few hours to several weeks.
Long-term storage methods
Storing the brain in formaldehyde for the long-term prior to use is an economical and convenient way to prevent microbial and autolytic degradation. It is especially convenient for gross tissue preservation for surgical training, as was performed in Alvernia et al. [
3] and Benet et al. [
9]. However, for histology purposes, storage in formaldehyde has been found to lead to a decrease in antigenicity over time. Lyck et al. [
58], who used this storage method, performed a quantitative study of several antigens over time, showing that antibody staining quality decreased for certain sensitive antigens, such as NeuN and CNPase, when stored in fixative over time. Similarly, McGeer et al. [
62] noted that brains fixed in formalin for a long period of time had negative staining results for the protein that they were studying, HLA-DR.
An alternative method for long-term storage for subsequent histology is to store tissues at sub-zero temperatures. However, this method requires the distribution of cryoprotectant throughout the tissue to prevent ice damage. Four studies reported using this method for long-term storage (Table
6). Notably, the glycerol-dimethylsulfoxide cryoprotectant method used by Insausti et al. [
42] has been found in non-human primate brain tissue to cause less tissue shrinkage than the sucrose-based methods [
27].
Table 6
Strategies for long-term storage of perfusion-fixed brain tissue
| Immersion in fixative | Surgical training | Separated head | 10% Formalin and 10% ethyl alcohol | 4 °C | Up to 4 years |
| Immersion in fixative | Surgical training | Separated head | 10% formaldehyde or 10% custom solution (ethanol 62.4%, glycerol 17%, phenol 10.2%, formaldehyde 2.3%, and water 8.1%) | 5 °C | Up to a year |
| Cryoprotection and freezing | Histology | 1 cm-thick coronal tissue blocks | Solutions of 10 and 20% glycerol in 0.1 M phosphate buffer and 2% dimethylsulfoxide | −80 °C | NR |
| Immersion in fixative | Histology | Whole brain | 0.1% paraformaldehyde in 0.15 M Sørensens phosphate buffer (pH 7.4) | 4 °C | Up to 4 years |
| Cryoprotection and freezing | Histology | 1 cm-thick coronal tissue blocks | Buffered 5% sucrose | −80 °C | NR |
| Cryoprotection and freezing | Histology | Tissue blocks (many 1 cm-thick) | 20–30% sucrose in 0.1 M phosphate buffer with 0.1% sodium azide | −80 °C | NR |
| Cryoprotection and freezing | Histology | Brain sections | 1.1 M sucrose, 37.5% ethylene glycol in PBS | −20 °C | NR |
To summarize, fixed brain tissue can be stored in fixative at refrigerator temperatures near 4 °C, but this will likely lead to a decrease in antigenicity over time. An alternative approach, which may allow for the preservation of antigenicity for longer, is to add cryoprotectant to the fixed brain tissue and store it at a freezer temperature such as − 80 °C.