Biomaterials 297 (2023) 122102
Contents lists available at ScienceDirect
Biomaterials
journal homepage: www.elsevier.com/locate/biomaterials
Activation of inflammasomes and their effects on neuroinflammation at the
microelectrode-tissue interface in intracortical implants
Melissa E. Franklin a, Cassie Bennett a, Maelle Arboite a, Anabel Alvarez-Ciara a,
Natalie Corrales a, Jennifer Verdelus a, W. Dalton Dietrich a, b, c, Robert W. Keane c, d, e,
Juan Pablo de Rivero Vaccari c, d, e, Abhishek Prasad a, c, *
a
Department of Biomedical Engineering, University of Miami, Miami, FL, USA
Department of Neurological Surgery, University of Miami Miller School of Medicine, Miami, FL, USA
The Miami Project to Cure Paralysis, University of Miami, Miami, FL, USA
d
Department of Physiology and Biophysics, University of Miami Miller School of Medicine, Miami, FL, USA
e
Center for Cognitive Neuroscience and Aging University of Miami Miller School of Medicine, Miami, FL, USA
b
c
A R T I C L E I N F O
A B S T R A C T
Keywords:
Inflammasome
Caspase-1
Utah electrode array
Foreign body response
Innate immune response
Invasive neuroprosthetics rely on microelectrodes (MEs) to record or stimulate the activity of large neuron as­
semblies. However, MEs are subjected to tissue reactivity in the central nervous system (CNS) due to the foreign
body response (FBR) that contribute to chronic neuroinflammation and ultimately result in ME failure. An
endogenous, acute set of mechanisms responsible for the recognition and targeting of foreign objects, called the
innate immune response, immediately follows the ME implant-induced trauma. Inflammasomes are multiprotein
structures that play a critical role in the initiation of an innate immune response following CNS injuries. The
activation of inflammasomes facilitates a range of innate immune response cascades and results in neuro­
inflammation and programmed cell death. Despite our current understanding of inflammasomes, their roles in
the context of neural device implantation remain unknown. In this study, we implanted a non-functional Utah
electrode array (UEA) into the rat somatosensory cortex and studied the inflammasome signaling and the cor­
responding downstream effects on inflammatory cytokine expression and the inflammasome-mediated cell death
mechanism of pyroptosis. Our results not only demonstrate the continuous activation of inflammasomes and
their contribution to neuroinflammation at the electrode-tissue interface but also reveal the therapeutic potential
of targeting inflammasomes to attenuate the FBR in invasive neuroprosthetics.
1. Introduction
Neuroprosthetic devices have marked capabilities to improve the
quality of life for individuals faced with neural injuries or neurodegen­
erative diseases such as spinal cord injury, stroke, paralysis, amyo­
trophic lateral sclerosis (ALS), Parkinson’s disease, and other
neuromotor disorders [1–4]. Electrodes are integral to neuroprosthetic
device design as tools for obtaining and modulating neuronal activity.
Accordingly, such devices are dependent on the ability to record or
modulate the activity of large assemblies of neurons via microelectrodes
(MEs). However, successful clinical integration of neuroprosthetics is
hindered since MEs have a critical limitation in reliably acquiring
neuronal signals for long durations [5–9], which can be attributed to
both material design and the foreign body response (FBR) that occurs at
the electrode-tissue interface. In recent years, Utah electrode arrays
(UEAs) have been applied in multiple human clinical trials [10–15],
however, studies that depict the evolving FBR to this array type are few
and limited [16–20].
Current research has revealed several cell types following ME im­
plantation that display a distinct immune response in the central ner­
vous system (CNS) and result in persistent neuroinflammation. These
cellular responses include, but are not limited to, astrocytes and
microglia activation leading to glial scarring [19,21–24], oligodendro­
cyte deformation [25,26], pericytes and endothelium reactivity [17], a
breach of the blood-brain-barrier (BBB) [17,18,27,28] and gliovascular
inflammation [25,26,29,30], and inflamed neurons undergoing
* Corresponding author. Department of Biomedical Engineering, The Miami Project to Cure Paralysis, University of Miami, 1095 NW 14th Terrace, Miami, FL,
33136, USA.
E-mail address: a.prasad@miami.edu (A. Prasad).
https://doi.org/10.1016/j.biomaterials.2023.122102
Received 11 October 2022; Received in revised form 16 March 2023; Accepted 23 March 2023
Available online 28 March 2023
0142-9612/© 2023 Elsevier Ltd. All rights reserved.
M.E. Franklin et al.
Biomaterials 297 (2023) 122102
degeneration [31,32]. The foreign body response exhibited from
implant-induced trauma is complex with various pathways that
contribute to overarching neuroinflammation. Recent work from our lab
shows the role of the complement cascade following ME implant injury
and elucidates the significance of an invoked innate immune system
following UEA implantation [16]. However, complement is only one
aspect of the intricate innate immune system that is affected due to an
injury to the CNS tissue. Synchronous crosstalk between complement
and other innate immune system components occurs to generate an
immune response [33–36]. An area of the innate immune system that is
stimulated under neuroinflammatory conditions [37–39] but has not
been well defined in context of ME implantation injury is the activation
of inflammasomes and their repercussions. The goal of this study is to
evaluate inflammasomes and reveal their importance within the innate
immune response towards neuroinflammation at the electrode-tissue
interface.
Inflammasomes are intracellular multiprotein complexes that
contribute to the initialization of the innate immune response [40,41].
Inflammasomes reside in the cytosol of stimulated cell types, predomi­
nantly resident perivascular macrophages [42], microglia [43], astro­
cytes [44], and neurons [45,46], and facilitate pro-inflammatory
caspase expression [41]. Research focused on defining the function of
inflammasomes has uncovered that injury to CNS tissue facilitate
inflammasomal sensor activation and thus inflammasome complex for­
mation [47–50]. In turn, inflammasome complexes evoke key inflam­
matory responses driven by activating caspase enzymes, primarily
caspase-1 46. Following caspase-1 activation, cleavage and release of
pro-inflammatory cytokines, like IL-1β and IL-18, cause a robust in­
flammatory response. Considered as the gatekeepers to highly damaging
neuroinflammation, activated interleukin family cytokines can induce a
form of dysfunctional cell lysis, known as pyroptosis, in both host and
surrounding cells [51]. Similar responses can be evoked solely from
caspase-1 activity [46]. Pyroptosis is a form of programmed cell death
characterized by dysregulated cytokine upregulation and a porous
cellular membrane rupture. Importantly, these destructive processes can
ultimately cause neuronal death [51]. Therefore, inflammasomes are
considered to be triggering, upstream proteins serving as directors in an
immune pathway and are suggested to drive pathogenesis within the
CNS resulting from acute infections, chronic diseases, and traumatic
injuries [41,52].
Fig. 1 illustrates the inflammasome signaling pathway which results
in pyroptosis caused by injury to the CNS tissue, such as ME implanta­
tion. The structure of a typical inflammasome complex consists of three
major components: 1) a cytosolic sensor, 2) an adaptor protein, and 3) a
bound caspase-1 enzyme [53]. Inflammasome sensors survey and
recognize foreign stimuli entering or released from the cellular micro­
environment, such as those derived from damage-associated molecular
patterns (DAMPs) or pathogen-associated molecular patterns (PAMPs)
pathways [54,55]. Upon recognition of DAMPs/PAMPs, a particular
inflammasome sensor oligomerizes and often recruits the adaptor pro­
tein apoptosis-associated speck-like protein containing a caspase
recruitment domain (ASC) [56]. Then, activated ASC binds to
pro-caspase-1 via CARD-CARD protein domain interactions, which
promotes maturation of the caspase-1 enzyme. Recruitment of active
caspase-1 cleaves inflammasome-mediated pro-inflammatory cytokines,
IL-1β and IL-18 [41,49,50,57,58], as well as membrane rupturing
enzyme, gasdermin-D, in turn facilitating the expression of these
markers and subsequent pyroptosis [59–61]. Different inflammasome
sensors have unique molecular compositions, receptive to varying,
corresponding stimuli [54]. For example, one of the inflammasome
sensors monitored throughout this study, NOD-like receptor protein-3
(NLRP3), has three subparts enabling its formation into a multiprotein
Fig. 1. Schematic of inflammasome cascade at the electrode-tissue interface. An overview of the innate immune pathway evaluated in this study. Upon ME
insertion, neural cells (magenta) at the interface including astrocytes (pink) and microglia (green) are stimulated and shift to activated phenotypes. Upon activation,
inflammasomal sensor proteins oligomerize and recruit ASC adaptor proteins which in turn recruit caspase-1 precursor enzymes. This three-part active multiprotein
structure is known as an inflammasome complex. Once intact, the inflammasome complex drives caspase-1 maturation. This results in interleukin-1 beta (IL-1β) and
interleukin-18 (IL-18) cytokines maturation as well as activating cleavage of gasdermin-D (GSDMD) membrane pore induction enzyme. Together, the inflammasomedependent pro-inflammatory cytokines and progressive membrane porosity are indicative of programmed cell lysis known as pyroptosis. (For interpretation of the
references to color in this figure legend, the reader is referred to the Web version of this article.)
2
M.E. Franklin et al.
Biomaterials 297 (2023) 122102
complex. The first subpart of the sensor’s structure is a protein-protein
interaction domain, second an oligomerization domain, and lastly a
putative sensor [53], which is represented as described in Fig. 1.
Our central hypothesis is that, as critical initiators of the innate
immune response, inflammasome complexes become activated
following microelectrode implant induced injury and act as mediators of
pyroptosis at the electrode-tissue interface. Unlike adaptive immunity
and secondary responses, the innate immune responses act quickly
following injury, which is the rationale for studying inflammasome
activation and roles across acute (48-hour, 72-hour), sub-acute (1-week,
2-week), and early chronic (4-week) time periods post UEA implanta­
tion. In this study, we report the activation and continued expression of
inflammasomes at the electrode-tissue interface at acute, sub-acute, and
early chronic periods following ME implantation, revealing their upre­
gulation and highlighting their critical role in neuroinflammation [62,
63] in intracortical UEA implants.
2.3. Quantitative real-time PCR (qRT-PCR)
At their respective timepoint, animals were anesthetized with iso­
flurane and given an overdose of a ketamine (150 mg/kg)-xylazine (20
mg/kg) cocktail followed by decapitation for euthanasia. For fresh tissue
harvest, the head cap and UEA were immediately and carefully removed
from the tissue and the brain tissue was gently rinsed in RNA-free water.
A 4 mm × 4 mm x 2 mm portion of cortical tissue where the electrode
resided was quickly extracted, weighed, flash frozen in liquid nitrogen,
and stored at − 80 ◦ C for future use in quantitative real time polymerase
chain reaction (qRT-PCR) protocols. From the time of euthanasia,
cortical samples were flash frozen within approximately 15 min to
prevent excessive RNA degradation [35,64]. For qRT-PCR, frozen brain
tissue samples were homogenized with 1 ml of PureZol (Bio-Rad, CA) at
room temperature for the proteins to completely dissociate from the
nucleic acids in the lysate sample. Once the RNA was isolated and pu­
rified, the purity and concentration of the extracted RNA was measured
with a spectrophotometer (NanoDrop One, ND-1000, ThermosFisher
Scientific, MA). Reverse transcription was then performed on the RNA
sample using a Superscript First-Strand Synthesis System (Bio-Rad, CA)
to obtain cDNA, which was stored at − 20 ◦ C. Gene expression analysis
was performed on a PCR detection system (CFX96 Touch, Bio-Rad, CA).
For each well on the plate, a PCR reaction was prepared containing 10 μL
Sybr-Green as the fluorescent label, 3 μL cDNA, 1 μL of the respective
gene primer, and 6 μL sterile water. Each gene primer was run in
duplicate wells. GAPDH was used as the reference gene. To monitor its
expression across PCR plates and ensure its suitability as a stable
reference gene, an average Cq value for GAPDH was calculated and
compared between unoperated control and experimental animal groups.
There was no statistical difference found between the groups which
indicates that GAPDH expression was stable and provided a stable
reference gene [65]. A summary of the gene targets examined in this
study, which represents the inflammasome activation and respective
downstream pathway, is listed in Table 1.
mRNA expression of each gene target (experimental) is represented
as fold-change values normalized to that of unoperated (control) tissue
samples read on the same well-plate. Both the experimental and control
sample Cq values were normalized to the reference gene, GAPDH Cq
value, producing ΔCq (ΔCq = Cq(target gene) - Cq(reference gene)). The
ΔΔCq value for each target gene was then calculated by normalizing
each experimental ΔCq value to the unoperated, control sample ΔCq
value (ΔΔCq = ΔCq(target gene) - ΔCq(control sample)). The relative
fold-change in mRNA expression was then calculated using the 2(− ΔΔCq)
method [66]. As relative fold-change in mRNA expression was normal­
ized to unoperated control samples, a fold-change of ≈1 was considered
similar to baseline physiological conditions while a 2-fold or larger
change in gene expression was considered significantly different
compared to controls [16,66]. Relative fold-change in gene expression is
presented as Mean ± SEM from each animal group with a dotted line
shown at 2-fold to indicate significant change in gene expression relative
to baseline expression in unoperated controls. An outlier ROUT test (Q
2. Methods
2.1. Overview and animal groups
All procedures were approved by the University of Miami Institu­
tional Animal Care and Use Committee. A total of 36 (n = 30 rats for
qRT-PCR and n = 6 rats for IHC) adult male Sprague-Dawley rats all
weighing approximately between 250 and 300 g were used in this study.
Animals were implanted with a custom-made, non-functional Utah
microelectrode array (UEA), with no tethering wires, into their so­
matosensory cortex. Each UEA consisted of 16-channels in a 4 × 4
configuration, 400 μm apart, of 1 mm long parylene coated silicon
shanks. Gene expression, quantified by qRT-PCR, was performed at
acute (48-hour, 72-hour), sub-acute (1-week, 2-week), and early chronic
(4-week) periods post-implantation (n=5 animals/group at each time­
point). Additionally, baseline gene expression was obtained from naïve
control animals (n = 5), which did not undergo surgery. For histological
assessments, immunohistochemistry (IHC) and FAM-FLICA Caspase-1
fluorescent assay was used to quantify protein expression at electrode
sites at the sub-acute (1-week) and early chronic (4-week) timepoints (n
= 3 animals/group).
2.2. Surgical procedure
Aseptic techniques were followed for all implant surgeries described
in detail previously [16–18]. Stereotactic surgery was performed to
implant non-functional UEAs into the rat cortex. All UEAs were gas
(ethylene oxide) sterilized and allowed to outgas at least 24-h prior to
starting the animal surgeries. Animals were anesthetized with isoflurane
and deep anesthesia (2% isoflurane, 1% oxygen) was maintained
throughout the duration of surgery. Following a midline incision, the
rat’s skull was exposed to locate the bregma and craniotomy location
was marked corresponding to the somatosensory cortex (1 mm lateral
and 2 mm posterior relative to the bregma). Four stainless steel screws
(3/16 inch, 0–80) were manually drilled into the skull for the headcap
assembly. A high-speed drill was used to make a craniotomy at the
marked location to expose the cortex. Upon resecting the dura, the
non-functional, untethered, floating UEA was gently placed on the
cortical surface and a pneumatic inserter (Blackrock Neurotech Inc, UT)
stereotactically positioned above the array was used to insert the array
into the cortex. Once the array was implanted, a thin, sterile 25 μm-thick
silastic sheet cut to a size slightly larger than the craniotomy, was used
to cover the craniotomy. Dental acrylic was then used to cover the entire
exposed skull surface. Following surgery, all animals were closely
monitored while they recovered on a heated recovering pad. An anal­
gesic (Carprofen, 5 mg/kg) was administered only on the day of surgery
and no other medications were used post-surgery.
Table 1
All gene primers for this study were purchased from Bio-Rad, CA. Unique Assay
IDs of each gene primer are included in Supplementary Table 1.
3
Abbreviations
Gene
NLRP1
NLRP3
NLRC4
AIM2
ASC (PYCARD)
CASP1
IL-1β
IL-18
GSDMD
GAPDH
NLR Family Pyrin Domain Containing 1
NLR Family Pyrin Domain Containing 3
NLR Family CARD Domain Containing 4
Absent in Melanoma 2
Apoptosis-associated speck-like protein containing a CARD
Caspase-1
Interleukin-1 beta
Interleukin-18
Gasdermin-D
Glyceraldehyde-3-phosphate dehydrogenase
M.E. Franklin et al.
Biomaterials 297 (2023) 122102
= 10%) was performed to identify any outlier value for each gene
transcript fold-change for each timepoint [67–70]. Then, a Shapiro-Wilk
test for normality was applied. If values displayed a normal distribution,
a one-way analysis of variance (ANOVA) was performed to determine
whether there was a significant difference in the fold change values for
each gene across time. If there was a significant difference (p < 0.05)
between the groups, a Tukey post-hoc test with correction was applied to
account for multiple comparisons. In the case of failing normality
testing, non-parametric Kruskal-Wallis testing was applied (p < 0.05)
followed by post-hoc Dunn’s test.
Reconstituted FAM-FLICA reagent was diluted in 1X PBS at 1:5 con­
centration and stored at − 20 ◦ C. Fixed tissue slides were serially washed
with 1X PBS before applying FAM-FLICA working solution (1:50) for 90
min in a dark room. Slides were then serially washed with 1X Apoptosis
Wash Buffer before applying coverslips. Once cured, slides were
immediately imaged to avoid fading of the fluorescent signal.
2.6. Imaging and analysis
Microscopy imaging was performed on a fluorescent microscope
(Eclipse Ti Series, Nikon Instruments, NY) and image analysis was per­
formed using ImageJ (NIH, MD). All images were captured at the same
exposure and gain for the corresponding immunohistochemical marker.
Images were captured at 40X magnification centering the site of elec­
trode injury, visualized as a “hole” in the image that represents where
the electrode shank was present. For each antibody and for the FAMFLICA histochemical probe, five electrode shank sites within a tissue
section labeled with the respective antibody or histochemical marker
and distributed across the array were used for quantification from each
animal. Images from each injury site (hole) for each immunohisto­
chemical marker and for all animals as well as the entire UEA footprint
for all animals are included in Supplementary Figs. 1–6. We and others
have reported the variability in obtaining electrode injury holes from
whole brain samples when explanting the UEA [16,19]. This has been
reported due to tissue adherence to the UEA shanks and cavitation due
to the array at the implantation site. Five injury sites that did not display
artifacts such as those occurring due to tissue tears and cavitation were
imaged for analysis.
Additionally, an image was captured at the same magnification and
equivalent image area (1.41× 105 μm [2]) from the contralateral (con­
trol) brain hemisphere, across from the electrode injury, from each an­
imal and for each marker to serve as a reference for normalization in
image analysis. Images were first converted to binary grey scale. For
each animal and histochemical marker, the background fluorescence
intensity was calculated as the mean grey value of all pixels over the
total area of the image (1.41 × 105 μm [2]) from the contralateral
control hemisphere image. Images from the injury site were normalized
with respect to the control contralateral brain tissue by subtracting the
background fluorescence for each target protein. An Otsu thresholding
filter was then applied to the images. The Otsu method [75,76] chooses a
threshold value that minimizes the intraclass variance of the thresh­
olded foreground and background or the black and white pixels. All
images were manually inspected after applying the thresholding algo­
rithm. Cellular count and area fraction metrics within each ipsilateral
image were then used to quantify target protein expression [77–84].
Following image quantification, any outlier area fraction or cell count
for each animal was removed using ROUT test for outliers (Q = 10%) to
account for potential imaging artifact and ensure a representative
averaged value for each timepoint subgroup [85,86]. The average area
fraction or cell count metric for each animal was calculated and the
values were then grouped by timepoint. A Shapiro-Wilk test of normality
2.4. Immunohistochemistry (IHC)
For immunohistochemistry (IHC), animals in the IHC groups were
euthanized via cardiac perfusions in order to prepare tissue extracts for
analysis. Briefly, deeply anesthetized animals were transcardially
perfused with 1X phosphate-buffered saline (PBS) followed by 4%
paraformaldehyde (PFA) to fix the tissue. Brain tissue was dissected and
cryoprotected in 30% (w/v) sucrose and stored at 4 ◦ C. For tissue
embedding, specimens were embedded in M1-embedding matrix
(Thermo Fisher Scientific, MA), and embedded molds were placed on
dry ice. The embedding mold containing the specimen was briefly sub­
merged in isopentane cooled over dry ice (<-65 ◦ C) until there were no
observed air bubbles releasing from the mold, to minimize freezing
artifact. 20 μm thick horizontal sections were cut on a cryostat (CM1520,
Leica Biosystems, IL) and stored at − 20 ◦ C for IHC. Brain tissue was
sectioned in horizontal slices to enable visualization of both the site of
electrode injury, referred to as ipsilateral indicating electrode arrayinduced injury, and the corresponding non-injured contralateral hemi­
sphere which served as the internal control for IHC analysis. A summary
of all primary and corresponding secondary antibodies used in this study
is presented in Table 2. For IHC, slides were first acclimated to room
temperature for 30 min and then placed in a blocking buffer solution
overnight at 4 ◦ C. Following blocking, respective primary antibody so­
lution was applied on the slides and incubated overnight at 4 ◦ C. The
slides were then subjected to serial washes with 1X PBS with 0.1%
Triton-X and then incubated in the secondary antibody solution at room
temperature for 2-hr. After the slides were washed and dried, Prolong
Diamond (Thermo Fisher Scientific, MA) was applied, and the slides
were cured in the dark at room temperature for 24-hr.
2.5. FAM-FLICA Caspase-1 histochemical assay
To determine active caspase-1 expression as a direct measure of
pyroptosis [72–74], a fluorescent labeled inhibitor of caspases (FLICA)
assay kit (Immunochemistry Technologies, Minneapolis, MN) was used.
The FLICA assay employs a green fluorescent inhibitor probe
FAM-YVAD-FMK to bind to caspase-1 specifically and irreversibly in
living cells [72–74]. As the probe becomes covalently coupled to the
caspase-1 enzyme, it is retained within an expressing cell whereas un­
bound FAM-FLICA diffuses out of the cell and is washed away.
Table 2
Summary of solutions used for immunohistochemical staining. The source for Ready-to-Use (RTU) Animal-Free Blocking Buffer (#5035100) was Vector Laboratories
(Burlingame, CA), while Normal Goat Serum (NGS) (#51097Z), Goat anti-mouse Cross Adsorbed Alexa 488 (#A32723) and Goat anti-rabbit Cross Adsorbed Alexa 555
(#A32732) secondary antibodies were purchased from Thermo Fisher Scientific.
Antibody
Supplier
Blocking Buffer Solution
Antibody
Classification
Species
Primary Antibody
Concentration
Secondary Antibody
Anti-ASC
As described in Desu [71] et al.
Polyclonal
Rabbit
1:250
AntiGFAP
Anti-Iba1
Thermo Fisher Scientific (MA512023)
FUJIFILM Wako Chemicals
(019–19741)
Millipore-Sigma (MAB3777)
Triton-X (0.4%)
RTU Animal-Free Blocking Buffer
Triton-X (0.4%)
RTU Animal-Free Blocking Buffer
10% NGS, Triton-X (0.2%) in 1x
PBS
10% NGS, Triton-X (0.2%) in 1x
PBS
Monoclonal
Mouse
1:1000
Polyclonal
Rabbit
1:1000
Monoclonal
Mouse
1:1000
Goat anti-rabbit, IgG
555
Goat anti-mouse, IgG
488
Goat anti-rabbit, IgG
555
Goat anti-mouse, IgG
488
AntiNeuN
4
M.E. Franklin et al.
Biomaterials 297 (2023) 122102
was used to determine whether the data was normal. An F-test was then
performed, and no differences were found between the variance be­
tween the groups (1-wk and 4-wk timepoints) for each immunohisto­
chemical marker. An unpaired, two-tailed, Student’s t-test was then
applied to determine whether there were any differences in the protein
expression between timepoints (1-wk and 4-wk). The data are reported
as average expressions of an immunohistochemical marker within a
standardized area in terms of Mean ± SEM and normalized to control
contralateral brain tissue.
expression of the key inflammasome sensor molecules and complexes.
The activation of sensor molecules and complexes leads to downstream
inflammasome-mediated pro-inflammatory signaling, which ultimately
results in inflammasome driven pyroptosis. The results are organized in
a sequential order that matches the initiation of inflammasome
signaling, its continuation, and termination. We first demonstrate the
activation of cell types that produce inflammasomal components. We
next reveal findings on inflammasome sensor molecule upregulation and
inflammasome complex formation. Then, we show the outcomes of
inflammasome activity with evidence of inflammasome-dependent
cytokine expression and pyroptosis.
3. Results and discussion
Activation of the inflammasome pathway is quantified by the
Fig. 2. Astrocytes and microglia as sources of inflammasomes: Glial cell activation at the site of electrode implant injury that are the major producers of
inflammasomes. GFAP was applied for labeling astrocytes and Iba1 for labeling microglia and macrophages. A, B) Images of an electrode injury site and of
contralateral, uninjured control tissue from each animal at 1-wk and 4-wk post-implantation are shown, labeled with GFAP (A) and Iba1 (B). All images used in the
analysis are included in Fig. 2 and 3 in the Supplementary document. Images were taken at 40x magnification and a 100 μm scale bar is displayed in the bottom right
corner. An average (Mean ± SEM) of the target protein positive area for each timepoint was calculated from IHC images for both C) GFAP and D) Iba1 expression.
The average positive area for the expressed protein was calculated using five distinct injury sites per animal (n = 3 animals/timepoint).
5
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Biomaterials 297 (2023) 122102
3.1. Role of astrocytes and microglia in inflammasome activation
melanoma 2 receptor (AIM2). NLRs and AIM2 are considered inflam­
masomal sensor molecules since they specifically drive the inflamma­
some activation pathway. Here, we evaluated the mRNA expression of
the sensor molecules that included NLRP3, NLRP1 (NLRP1a), NLRC4,
and AIM2 at various acute and sub-acute timepoints. Each of these
inflammasome receptors, with unique molecular structures, dictate
inflammasome activation pathways that occur within different neural
cell populations [47,48]. NLR pyrin domain containing 3 (NLRP3) is the
most studied and thought to be the most abundant inflammasome sensor
receptor [48] which is mainly found in microglia and astrocytes [48,88,
101,102]. NLRP1 was originally discovered as a sensor expressed in
neurons [45,56] and is shown to be critically involved in several CNS
injury models [56,62,103,104]. Specifically, NLRP1a expression was
evaluated in our study since this is the homolog sequence of the NLRP1
gene prominently expressed within the rat genome. Further, NLRP1a is
the ortholog sequence of the NLRP1 gene detected within the human
genome, making it ideal to study for clinical relevance [105–107]. NLR
caspase recruitment domain (CARD) containing 4 (NLRC4) does not
require adaptor priming to induce procaspase-1 108; however, it is sug­
gested to increase cytokine expression upon binding of ASC protein and
is involved in inflammation within glial cell types [108,109]. Further­
more, AIM2, which requires ASC adaptor bridging similarly as to NLRP3
and NLRP1 [110], is hypothesized to regulate neuronal pyroptotic death
[51].
There was significant upregulation (>2-fold change relative to
unoperated control animals) in the mRNA expression of all sensor
molecules by 72-hr post-implant, indicating the activation of
inflammasome-mediated pathways at these early timepoints following
injury (Fig. 3). At 48-hr post-implant, NLRP1 and NLRP3 had modest
upregulation in mRNA expression (~2-fold change), which further
increased at 72-hr (~4-folds) and remained upregulated throughout the
4-wk period (Fig. 3A and B). The mRNA expression of NLRP1 remained
consistently elevated for all the timepoints tested (Fig. 3A). The mRNA
expression of NLRP3 depicted temporal changes in the expression across
the tested timepoints. NLRP3 expression peaked at 1-wk (~4 foldchange, 48-hr vs. 1-wk **p = 0.0095) and decreased by 2-wk (1-wk
vs. 2-wk ***p = 0.003) (Fig. 3B). AIM2 also displayed greatest magni­
tude of fold-change relative to unoperated controls at 1-wk (~5-fold
change) but returned to control values (<2-fold change) by 4-wk
(Fig. 3C). Additionally, NLRC4, similar to NLRP1 and NLRP3, became
significantly upregulated compared to unoperated controls (>2-fold) by
48-hr. NLRC4 remained consistently elevated throughout the 2-wk
period (~6 fold-change) before declining by 4-wk (~2 fold-change)
(Fig. 3D). A summary of all inflammasome sensor molecules evaluated
was plotted along the same y-axis and organized by timepoint to show
the differences in fold-change magnitudes between gene transcripts and
to visualize the concurrent temporal patterns of all molecules simulta­
neously (Fig. 3E).
These results indicate the activation of inflammasome sensor mole­
cules following electrode implant injury remain elevated during the
acute and early-chronic timepoints tested in this study. Their activation
even at the earliest timepoints (48-hr) indicate the inflammasome acti­
vation occurs during the acute period following injury, while their
consistent elevation several weeks following implant-induced injury
suggests the ongoing, persistent neuroinflammation due to the presence
of a foreign body in the brain tissue. Thus, this study begins to define
which inflammasome sensors are dominant within UEA implantation
injury model. Both inflammasome sensor molecules, NLRP1 and NLRP3,
upregulated by 48-hr and remained distinctly elevated at 4-wks
(Fig. 3E). This suggests that these two sensors may play vital roles in
activation of inflammasome complexes during the acute and sub-chronic
periods following electrode implant-induced injury. These results are of
interest as NLRP1 is more prominently released in neurons whereas
NLRP3 is primarily expressed in supporting glial populations [41]. Our
results indicate NLRP1 is significantly elevated at 4-wk timepoint
compared to NLRP3, AIM2, and NLRC4 (***p = 0.002, ****p < 0.0001,
Within the CNS, microglia and astrocytes are considered the primary
effectors of neuroinflammation [87]. Microglia act as first responders
when conditions stray from homeostasis and when there are insults to
the CNS [38]. Once triggered, microglia become activated and display a
phenotypic shift, potentially assuming various phenotypes including
phagocytic macrophages [38]. Throughout this activation, microglia
and macrophages host and secrete a variety of pro-inflammatory me­
diators, including inflammasome complex components and subse­
quently, matured cytokines [38,43,88]. Similar to activated microglia
and infiltrated perivascular macrophages, astrocytes are also involved in
immune defense. Upon injury to the CNS, this cell population undergoes
hypertrophy and hyperplasia—a process known as astrogliosis [38].
Throughout early astrogliosis, astrocytes form long filaments to isolate
the injury, developing an astrocytic scar that can protect neural net­
works from progressive damage [89–92]. However, under inflamed
conditions, astrocytes can secrete pro-inflammatory cytokines, where
persistent astrogliosis can prevent neuronal regeneration [38]. Addi­
tionally, cross-communication between microglia and astrocytes can
exacerbate the innate immune response. In fact, there is evidence of
initial microglial induction leading to astrocytic inflammasomal pro­
duction [93]. Therefore, it is important to assess the state of glial pop­
ulations in context with inflammasome activation throughout UEA
implantation due to their roles in the immune response signaling and as
the major producers of inflammasomes.
We assessed localized astrocytic (GFAP) and microglial/macro­
phagic (Iba1) expression at the 1-week and 4-weeks timepoints through
IHC (Fig. 2). Iba1 can detect both microglia and macrophages [94] and
thus, suitable for studying the inflammasome cascade since inflamma­
some components activate in both cell phenotypes [95]. GFAP and Iba1
were both abundantly expressed, and their expression levels remained
elevated through the early chronic-period of 4-weeks (Fig. 2), indicating
astrocytic and microglial activation at the electrode-tissue interface.
Although glial cell activation is well documented within the literature
following electrode implant injury [6,7,19,23,89,91], it is important to
demonstrate the activation of these cell types as they are the major
sources of inflammasomes [42–44,46,95]. Further, glial cell activation
in conjunction with inflammasome activation for neural implants is
unexplored and can provide insights into the mechanisms that trigger
the innate immune system and the secondary inflammatory cascades
after initial glial cell activation. Both astrocytes and microglia are
known to quickly react to DAMPs and PAMPs stimulation, resulting in
inflammasomal subparts expression. The glial cell activation, such as
evident in the electrode implant injury, is among the earliest events in
the
inflammatory
cascade
[23,30]
that
results
in
inflammasome-mediated downstream activity [38,96–98].
3.2. Activation of inflammasome sensor molecules
A panel of inflammasome sensor molecule gene transcripts were
monitored, as their expression facilitates the activation of the inflam­
masome complex [37,39,99]. As described previously, the structure of a
typical inflammasome complex consists of three major components: a
cytosolic sensor, an adaptor protein, and a bound caspase-1 enzyme
[53]. Once activated, these three-part sensor molecules bind to other
inflammasome subparts and are expressed in response to released
cytosolic DAMPs and/or PAMPs. DAMPs/PAMPs are considered initial
signals in response to danger stimuli introduced to the cellular micro­
environment. PAMPs are typically derived from external microorgan­
isms causing exogenous pathogen-based inflammation, whereas DAMPs
are sourced internally from cells undergoing different forms of endog­
enous distress, such as trauma or injury [54,55,100]. Once activated,
DAMPs/PAMPs bind to a class of molecules called pattern recognition
receptors (PRRs) to drive inflammatory pathways. Some PRRs include
toll-like receptors (TLRs), NOD-like receptors (NLRs), and absent in
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Biomaterials 297 (2023) 122102
Fig. 3. Gene expression of local inflammasome sensor molecules in UEA-implanted animals. Sensor molecules A) NLRP1, B) NLRP3, C) AIM2, and D) NLRC4
were quantified for their mRNA expression (Mean ± SEM) using qRT-PCR at 48-hr, 72-hr, 1-wk, 2-wk and 4-wk post-implantation surgery. Gene expression is
presented as fold changes relative to healthy, unoperated control animals, where 1-fold is considered a baseline value similar to healthy, unoperated controls. A
dashed line at 2-fold is to emphasize asignificant increase in gene expression relative to control, baseline expression. NLRP1 data includes n = 5 animals for each
timepoint. NLRP3 and AIM2 data includes n = 4 animals at 72-hr and 2-wk and n = 5 animals for 48-hr, 1-wk, and 4-wk. NLRC4 data includes n = 4 animals at 72-hr
and 4-wk and n = 5 animals for 48-hr, 1-wk, and 2-wk timepoints, after performing a ROUT test to remove outliers from the data sets. There were temporal changes
in NLRP3 gene expression across tested timepoints (48-hr vs. 1-wk **p = 0.0095), (72-hr vs. 2-wk **p = 0.0018), and (1-wk vs. 2-wk ***p = 0.003). E) Summary plot
of mRNA expression of all sensor molecules grouped by timepoint to compare their expression levels. A one-way ANOVA and post-hoc Tukey tests were performed at
each timepoint, comparing the fold-change values between the four sensor molecule gene transcripts. At 48-hr, NLRC4 was significantly elevated compared to
NLRP1, NLRP3, and AIM2 (***p = 0.0001, ***p = 0.0001, and ****p < 0.0001 respectively). At 2-wks, there was significant difference in expression between NLRP3
and NLRC4 (*p = 0.0175). At 4-wks, there was significant elevation of NLRP1 compared to NLRP3, AIM2, and NLRC4 (***p = 0.002, ****p < 0.0001, and ****p <
0.0001 respectively).
and ****p < 0.0001 respectively) (Fig. 3E). These results may be sug­
gestive of which cell populations activate specific sensor molecules, such
that neurons expressing NLRP1 are triggered later in the FBR in com­
parison to glial cells that are the predominant sources of the other sensor
molecules (Fig. 3E) [45,48,56,62,88,102,111–113]. Therefore, these
data indicate that in addition to inflammasome sensors being present,
they could differentially express across multicellular populations in
response to both acute and secondary injury from the presence of a
foreign body.
Despite the evidence in support of AIM2 expression contributing
towards long-term functional impairments such as in post-stroke [114,
115], this sensor molecule appears to be the least consistently elevated
throughout the timepoints following electrode array implantation. This
may be attributed to which DAMPs/PAMPs are released that are capable
of triggering AIM2 expression [51]. It is likely that AIM2 expression is
due to the presence of a microelectrode array as the foreign body in the
CNS tissue. Furthermore, the temporal pattern of NLRC4 activity is
interesting because it had robust elevation by 48-hr and the greatest
relative fold change at the 72-hr timepoint. Our results indicate NLRC4
is significantly elevated at 48-hr compared to NLRP1, NLRP3, and AIM2
(***p = 0.0001, ***p = 0.0001, and ****p < 0.0001 respectively).
Interestingly, NLRC4 does not require ASC binding protein to be func­
tional and can form an inflammasome complex independently [48,116,
108,110]. Thus, the data provide preliminary support that NLRC4’s
independent activity may contribute to earlier (by 48-hr) downstream
effects of inflammasome mediation such as proteolytic cleavage of
caspase-1 and cytokine generation in comparison to the other sensor
molecules evaluated (Fig. 3E). These findings provide strong support
that upstream inflammasome sensor molecules are present early
following UEA implantation and remain elevated to cause sustained
inflammasome-mediated neuroinflammation.
3.3. Formation of inflammasome complexes indicated by ASC specks and
caspase-1
Inflammasome sensor molecules rely on adaptor proteins, such as the
apoptosis associated speck-like proteins containing a caspase recruit­
ment domain (ASC), for the recruitment and activation of caspase-1
from its precursor form [117]. Once the inflammasome complex is
activated, ASC clusters into large protein assemblies, referred to as ASC
specks. Because ASC is a key measure of an intact inflammasome com­
plex, ASC expression levels have become a standard readout for
inflammasome complex activation [118]. In addition to the pyrin and
CARD domain (PYCARD) gene that encodes ASC, the caspase-1 gene was
also monitored via qRT-PCR due to its maturation post-inflammasome
complex formation. Further, the transcribed ASC protein specks were
measured to support PYCARD expression at two corresponding time­
points, 1-wk and 4-wk via IHC, and validate the measurement of intact,
aggregated inflammasome complex.
The gene encoding for ASC protein (PYCARD) was significantly
upregulated (~5-fold) by 48-hr and remained elevated throughout the
implant duration of 4-weeks (Fig. 4C). This consistent expression of the
PYCARD gene encoding for inflammasome complex activation was
validated by IHC of the transcribed ASC protein specks at 1-week and 47
M.E. Franklin et al.
Biomaterials 297 (2023) 122102
Fig. 4. Formation of inflammasome complexes indicated by ASC specks and caspase-1. A) Images of an electrode injury site and of contralateral, uninjured
control tissue from each animal at 1-wk and 4-wk post-implantation labeled with ASC are shown. All images used in the analysis are included in Fig. 4 in the
Supplementary document. In general, images reveal localized ASC specks surrounding ME injury sites. Images were taken at 40x magnification and a 100 μm scale
bar is displayed in the bottom right corner. B) An average (Mean ± SEM) of the target protein positive area for each timepoint was calculated from IHC images for
ASC expression. The average positive area for the expressed protein was calculated using five distinct injury sites per animal (n = 3 animals/timepoint). Corre­
sponding mRNA expression of the C) PYCARD gene encoding ASC and D) Caspase-1, using qRT-PCR, show elevated levels of inflammasome complex components
across all tested timepoints relative to unoperated controls (>2-fold). Data is presented as the Mean ± SEM for each timepoint. After applying an outlier test, both the
PYCARD and Caspase-1 qRT-PCR data includes n = 5 animals for 48-hr, 72-hr, 1-wk, and 2-wk and n = 4 animals for the 4-wk timepoint, respectively. There were
significant differences in caspase-1 expression between the 48-hr and 1-wk (*p = 0.0472) and 2-wk (*p = 0.0207) timepoints.
weeks. Animals (n = 3/group) displayed similar expression of the ASC
protein quantified as ASC-positive area of ASC specks (Mean ± SEM)
across the 1-wk (13800.20 ± 1783.31 μm2) and 4-wk (15232.40 ±
2666.27 μm2) timepoints (Fig. 4A and B). Moreover, ASC speck activity
was visually similar at both timepoints post-implantation. ASC speck
activity appears as localized clusters of specks around the injury site
(hole made by the electrode shank) and more distributed specks as
observed farther away (Fig. 4A). Additional evidence of inflammasome
complex activation is demonstrated by the consistent expression of
caspase-1 gene transcript (Fig. 4D). mRNA expression levels of caspase-1
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M.E. Franklin et al.
Biomaterials 297 (2023) 122102
were significantly elevated relative to controls and increased from ~2fold at 48-hr, to ~7-fold at 1-wk (*p = 0.0472) and at 2-wks (*p =
0.0207).
The oligomerization of ASC into specks provides a platform for cas­
pase recruitment [119]. Specifically, procaspase-1 is recruited through
the CARD-CARD interactions of ASC [120]. These results indicate suc­
cessful redistribution of ASC adaptor protein from its sporadic expres­
sion throughout the cellular cytosol into activated ASC speck clusters
[121]. ASC consists of two domains—a pyrin domain (PYD) and caspase
recruitment domain (CARD), with both playing roles in the formation of
an intact inflammasome complex [121]. Elevated and sustained
expression of PYCARD gene transcript as well as labeling of anti-ASC
speck protein indicates sustained expression of the activated, tran­
scribed inflammasome binding protein. The caspase-1 mRNA levels are
elevated compared to unoperated controls, suggesting there is gene
expression for the mature enzyme versus its inactive, precursor form.
Combining ASC and caspase-1 results show acute inflammasome com­
plex formation as well as sustained complex activation. These results
underscore the presence of inflammasome complex activation
throughout the implant duration tested and that there is evidence of a
sustained innate immune response following device implantation.
Fig. 5. Elevated expression of IL-1β and IL-18 levels suggestive of
inflammasome mediated cytokine activity. IL-1β and IL-18 serve as strong
downstream indicators of inflammasome-mediated cytokine expression as the
processing mechanism of both these pro-inflammatory cytokines is initiated
upon the formation of an inflammasome complex followed by cleavage of the
interleukin precursors by caspase-1, before they can be secreted by the host cell.
Sustained and elevated mRNA expression for A) IL-1β and B) IL-18 are sug­
gestive of inflammasome mediated pro-inflammatory cytokine expression. Data
is presented as the Mean ± SEM for each group. For IL-1β, the data includes n
= 4 animals at 48-hr, 72-hr, 1-wk, 2-wk, and n = 5 animals at 4-wk after
applying outlier test. For IL-18, data includes n = 4 animals for 4-wk timepoint
and n = 5 animals for all other timepoints. There were significant differences in
IL-1β fold-change values between 72-hr and 4-wk (*p = 0.0425).
3.4. Inflammasome activation mediates IL-1β and IL-18 expression
Cytokines IL-1β and IL-18 are critical mediators of neuro­
inflammatory and neurodegenerative disorders in both acute CNS injury
and chronic conditions such as multiple sclerosis (MS) [53] and Alz­
heimer’s [122,123]. IL-1β is rapidly upregulated following traumatic
brain injury and has shown increased expression throughout the onset of
neuro-pathologies [71,95,124]. We have previously shown upregulation
in the mRNA expression of IL-1β among several interleukin family gene
transcripts which take part in the pro-inflammatory signaling in the
acute period following intracortical implantation of UEA [17,18]. In this
study, we present the concurrent expression of interleukins and
inflammasome complexes in an electrode implant injury.
Despite conflicting evidence pointing to IL-1β′ s potential role in
neuroprotection and repair, the predominating view remains that IL-1β
is regarded as a potent initiator of neuroinflammation [125]. Similarly,
IL-18 is a key pro-inflammatory cytokine in the CNS, controlling two
distinct immunological regulatory pathways that induce cytotoxicity
and inflammation. First, IL-18 triggers matrix metalloproteinase (MMP)
expression and exacerbates expression of other pro-inflammatory cyto­
kines such as tumor necrosis factor (TNF) and IL-1β. Additionally, IL-18
is heavily expressed in glial cells including microglia and astrocytes and
facilitates expression of apoptotic factors within these cell populations
under inflamed conditions [122,126–129].
Both IL-1β and IL-18 have unconventional processing mechanisms
compared to most other cytokine proteins that are vesicle-packaged
through the cellular Golgi apparatus before extracellular secretion
[130,131]. This non-traditional cytokine processing initiates once the
three-part inflammasome complex is formed consisting of a (1) cytosolic
sensor consisting of three-subunits, (2) ASC, and (3) pro-caspase 1.
Then, mature caspase-1 becomes activated and proteolytically cleaves
the interleukin precursors from their pro-state into their mature forms
that are then to be secreted by the host cell [41]. Thus, measuring IL-1β
and
IL-18
serve
as
strong
downstream
indicators
of
inflammasome-mediated cytokine expression. Similar to other gene
transcripts involved in the inflammasome signaling evaluated in this
study, mRNA expression for IL-1β and IL-18 were significantly elevated
compared to control animals (no surgery) as early as 48-hr (Fig. 5). A
Kruskal-Wallis test was applied and post-hoc Dunn’s testing determined
significant differences in IL-1β fold-change values between 72-hr and
4-wk (*p = 0.0425). IL-1β was robustly upregulated (~15–30 fold
relative to unoperated controls) throughout the first week post-UEA
implantation before gradually declining by the 4-wk period (Fig. 5A).
IL-18 mRNA expression level, relative to unoperated controls, was also
found to be increased (~3-6-fold) in the first week following implan­
tation, which returned to baseline levels by the 2-wk timepoint (Fig. 5B).
Sustained elevation in the mRNA levels of IL-1β and IL-18 in
conjunction with the presence of ASC and PYCARD highlight the neu­
roinflammatory downstream molecules associated with inflammasomes
and indicate the inflammasome complex activation as early as 48-hr
following electrode implantation and continuing through the acute
and early chronic (4-wk) periods. First, matured caspase-1 activation is
regulated through inflammasome formation, which cleaves preformed
pro-IL-1β and pro-IL-18 during the initial “priming” stage. These data
display the “activation” stage in which mature cytokines are generated
[132]. It is hypothesized that this two-stage mechanism required to
induce cytokine expression allows for greater control, timing, and
magnitude of the inflammatory cascade [132]. Although it is important
to consider alternative routes for IL-1β/1L-18 cytokine production that
can contribute to neuroinflammation, including caspase-4, -5,-11 [133,
134] and caspase-8/NF-κB/Fas signaling pathways [135,136], inflam­
masome mediation and subsequent caspase-1 activity is regarded as a
key, classical mechanism for IL-1β/1L-18 upregulation [46,133,137].
Given both inflammasome-mediated caspase-1 activity’s role in IL-1β
induction and that IL-1β remains significantly upregulated by the
4-week timepoint, our results demonstrate that inflammasome activity
and persistent inflammation in the early chronic phase coexist. Previous
studies have revealed different regulatory functions of separate inter­
leukin types in a rat ischemic model [138,139]. It is of interest that
expression of both IL-1β and IL-18 peaks by 72-hr; yet IL-1β remains
elevated into the chronic period whereas IL-18 returns to homeostatic
levels. This may suggest differential roles that cytokines play in the in­
flammatory signaling which could explain their modulation for varying
durations following implantation injury. Another study within a mouse
neuroinflammatory model found IL-18 induces various other cytokine
and chemokine activity to a greater degree compared to IL-1β, and thus
hypothesized IL-18 to have more potent pro-inflammatory signaling
effects [140]. This could also be a potential explanation for the differing
chronic trends between IL-1β and IL-18 seen within an electrode im­
plantation model, such that IL-18 may serve as a major effector for
cytokine production in the earlier phases of injury. Further analysis
would be required to distinguish the distinct roles of these
inflammasome-mediated inflammatory cytokines following electrode
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Biomaterials 297 (2023) 122102
implantation.
However,
these
results
show
a
sustained
pro-inflammatory microenvironment that could be attributed to
inflammasome signaling. Moreover, combining these cytokine expres­
sion patterns with the inflammasomal sensor molecule temporal trends,
we can identify the inflammasomal cascades and downstream outcomes
on neuroinflammation as a result of ME implantation injury in the CNS.
Fig. 6. Caspase-1 mediated pyroptosis at the site of UEA implantation indicative of inflammasome driven cellular changes. A, B) Images of an electrode
injury site (ipsilateral) and of contralateral uninjured control tissue from each animal at 1-wk and 4-wk post-implantation are shown, labeled with FAM-FLICA-CASP1
(A) or NeuN (B). All images used in the analysis are included in Fig. 5 and 6 in the Supplementary document. Images were taken at 40x magnification and a 100 μm
scale bar is displayed in the bottom right corner. The average positive area for the expressed protein was calculated using five distinct injury sites per animal (n = 3
animals/timepoint). FAM-FLICA-CASP1 probe was applied to bind to the active caspase-1 enzyme, which is the expression of activated caspase-1 enzyme and thus,
indicative of caspase-1 mediated pyroptosis. FAM-FLICA-CASP1 signal was found to be concentrated at the electrode injury site at 1-wk, which became more
concentrated and distributed by 4-wks. In general, these results are indicative of increasing pyroptotic activity at the site of electrode implant injury. Additionally,
NeuN antibody was used to label neurons at the corresponding timepoints of 1-wk and 4-wks. Interestingly, an increase in caspase-1 driven pyroptosis coincided with
a declining trend in neuronal density. It is important to note that FAM-FLICA-CASP1 and NeuN were not co-labeled because of different IHC labeling protocols. C) An
average (Mean ± SEM) of the target protein area, across timepoints, for FAM-FLICIA-CASP1 expression shows significantly increased caspase-1 activity between the
timepoints (*p = 0.043), thus, providing evidence of pyroptosis occurring at injury sites. D) An average of the NeuN positive cells at 1-wk and 4-wk display a
discernible decrease in neurons over time coincident with an increase in pyroptotic activity at the injury site. E) Gene expression, shown as Mean ± SEM, indicate
elevated GSDMD expression post ME implantation relative to unoperated control animals, providing additional support that pyroptotic cell lysis occurs at the site of
electrode implantation (n = 5 animals/timepoint).
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Biomaterials 297 (2023) 122102
3.5. Inflammasome driven cellular changes: evidence of caspase mediated
pyroptosis
inflammasome components, we first showed the activation of glial cells
(macrophages, microglia, and astrocytes) at the site of electrode implant
injury. We next showed persistent elevation of the various components
of the inflammasome complex that includes inflammasome sensor
molecules, the ASC binding protein, and caspase-1 gene throughout the
4-wk implant duration. These results highlight that the three vital
components of inflammasome complexes are present and active at the
electrode injury site as a chronic problem. Next, we showed the effect of
inflammasome activation on downstream effector cytokines of IL-1β and
IL-18, the expression of which were upregulated soon after ME im­
plantation and remained elevated for the respective implant duration.
These results are indicative of the downstream outcomes of
inflammasome-mediated neuroinflammation since these cytokines can
rapidly exacerbate inflammation through recruitment of other cytokines
and chemokines as well as exert effects on surrounding cells. Finally, we
demonstrate how inflammasome activation induces pyroptosis, a
cellular process in which caspase-1 cleaves and activates effector cyto­
kines IL-1β and IL-18 and drives pyroptotic membrane degradation by
binding to gasdermin-D (GSDMD), a marker for cell lysis. The results
indicate sustained presence of GSDMD, providing further evidence of
elevated levels of pyroptosis occurring at the injury site. Interestingly,
pyroptotic activity at the injury site coincided with a decrease in
neuronal density. It is also important to note the limitations of the study
which included the inability to co-label FAM-FLICA and NeuN because
of different labeling protocols, lack of electrophysiological monitoring
due to non-functional UEAs, and the need for further experimental
validations to generalize the presented findings for other ME platforms
which have significantly distinct array footprint and method of inser­
tion. In summary, this study demonstrates the continuous activation of
inflammasomes at the electrode-tissue interface, inflammasomemediated neuroinflammation and pyroptosis that could potentially
lead to neuronal cell loss. Unlike a traumatic brain injury where the
injury occurs acutely, MEs reside in the CNS tissue, resulting in a
persistent neuroinflammation even at long chronic durations [7,19,20,
23,24]. Therefore, uncovering multiple mediators that contribute to
neuroinflammation will build a greater understanding of the intricate
immune response following ME insertion as well as serve to reveal po­
tential therapeutic targets to minimize neuroinflammation in response
to biomaterial integration within the CNS.
Caspase-1 enzyme plays a dual role in cellular inflammation and
subsequent cellular deconstruction [57,61]. Not only is caspase-1
responsible for the cleavage and activation of the effector cytokines
IL-1β and IL-18, but it also drives the membrane rupturing characteristic
of the programmed, inflammatory cellular death process of pyroptosis
[59,60,141]. Caspase-1 drives pyroptosis by binding to gasdermin-D
(GSDMD), which in turn forms pores at the cellular membrane [57,
59–61]. The increasing porosity at the cellular membrane leads to lysis
and leakage of the inflamed cell’s contents [47–51]. Importantly, this
includes the inflammatory cytokines of the host cell that are released
into the extracellular matrix and exacerbate inflammation within the
microenvironment by later affecting surrounding, proximal cells
[47–51]. Since IL-1β and IL-18 specific cytokine production and mem­
brane deformation are signs of inflammatory cell death triggered by
caspase-1, activity of the potent caspase-1 enzyme is considered a direct
measure of pyroptosis [61,72,73].
FAM-FLICA was applied to bind to the active caspase-1 enzyme,
inhibiting its further enzymatic activity from cytokine activation and/or
cell lysis progression, to quantify pyroptotic activity at the site of
electrode-implant injury. The FAM-FLICA assay to measure pyroptosis
was evaluated at 1-wk and 4-wk following UEA implantation. The
expression of the FAM-FLICA probe, which is the expression of activated
caspase-1 enzyme, significantly increased (*p = 0.043) from the 1-wk to
4-wk timepoint (Fig. 6A and C). This aligns with the aforementioned
finding of consistent upregulation of the caspase-1 gene transcript dur­
ing the 4-wk implant duration (Fig. 4D), which can ultimately lead to
accumulated protein translation and enzyme activity. At 1-wk, FAMFLICA signal was found to be concentrated at the electrode injury site
which became more concentrated and distributed by 4-wk. This in­
dicates that while caspase-1 activity begins soon after injury by 1-wk,
caspase-1 expression as assessed through FAM-FLICA increases signifi­
cantly (change of ~3-folds) as the implant duration progresses by 4-wk.
Consequently, these results are suggestive of increasing pyroptotic ac­
tivity at the site of electrode implant injury.
To further show evidence of pyroptosis, mRNA expression of GSDMD
was evaluated and found to be elevated at and beyond the 72-hr time­
point (~2.5-fold change) relative to unoperated control animals. As
GSDMD is a marker for cell lysis, results indicative of sustained upre­
gulation of GSDMD provides further evidence of elevated level of
pyroptosis occurring at the site of electrode implant injury (Fig. 6E).
Additionally, NeuN antibody was used to label neurons at the corre­
sponding timepoints of 1-wk and 4-wks. Interestingly, an increase in
caspase-1 driven pyroptosis coincided with a declining trend in neuronal
density (Fig. 6B and D). It is important to note that FAM-FLICA and
NeuN were not co-labeled because of different labeling protocols for the
immunohistochemical marker NeuN and the flurocolorometric histo­
logical probe FAM-FLICA. Although the study did not ascertain directly
that changes in neuronal densities were as a result of inflammasome
driven pyroptosis, our study provides direct evidence of caspase medi­
ated pyroptosis occurring concurrently with a reduction in neuronal
densities at the electrode implant site.
Credit author statement
MEF, Conceptualization, performed experiments, data analysis,
writing and editing the paper. CB, performed experiments and editing
the paper. MA, performed experiments, writing, and editing the paper.
AA, performed experiments. NC, JV, performed experiments, data
analysis, and editing the paper. WDD, RWK, conceptualization, data
visualization, editing the paper. JPdRV, AP, conceptualization, experi­
mental design, data analysis, writing and editing the paper, acquired
funding for the project.
Declaration of competing interest
The authors declare the following financial interests/personal re­
lationships which may be considered as potential competing interests:
JPdRV, RWK and WDD are co-founders and managing members of
InflamaCORE, LLC and have licensed patents on inflammasome proteins
as biomarkers of injury and disease as well as on targeting inflamma­
some proteins for therapeutic purposes. JPdRV, RWK and WDD are
Scientific Advisory Board Members of ZyVersa Therapeutics.
4. Conclusions
Within the central nervous system, inflammasomes play a critical
role in the initiation of the innate immune response and their activation
induces pyroptosis, which is a form of programmed cell death. The
objective of this study was to demonstrate inflammasome activation and
its continued presence at the site of electrode implant injury at acute,
sub-acute, and early chronic periods following UEA implantation,
highlighting their role in neuroinflammation and inflammasome medi­
ated cellular changes resulting in pyroptosis. Since immune cells are the
initial responders to a foreign body stimulus and are major sources of the
Data availability
The raw/processed data required to reproduce these findings cannot
be shared at this time as the data also forms part of an ongoing study.
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Biomaterials 297 (2023) 122102
Acknowledgements
[23] T.D. Kozai, et al., In vivo two-photon microscopy reveals immediate microglial
reaction to implantation of microelectrode through extension of processes,
J. Neural. Eng. 9 (2012), 066001. PMID: 23075490.
[24] D.H. Szarowski, et al., Brain responses to micro-machined silicon devices, Brain
Res. 983 (2003) 23–35. PMID: 12914963.
[25] S.M. Wellman, et al., The role of oligodendrocytes and their progenitors on neural
interface technology: a novel perspective on tissue regeneration and repair,
Biomaterials 183 (2018) 200–217. PMID: 30172245.
[26] K. Chen, et al., In vivo spatiotemporal patterns of oligodendrocyte and myelin
damage at the neural electrode interface, Biomaterials 268 (2021), 120526.
PMID: 33302121.
[27] J.D. Falcone, et al., Correlation of mRNA expression and signal variability in
chronic intracortical electrodes, Front. Bioeng. Biotechnol. 6 (2018) 26. PMID:
29637071.
[28] T. Saxena, et al., The impact of chronic blood-brain barrier breach on intracortical
electrode function, Biomaterials 34 (2013) 4703–4713. PMID: 23562053.
[29] T.D. Kozai, et al., Reduction of neurovascular damage resulting from
microelectrode insertion into the cerebral cortex using in vivo two-photon
mapping, J. Neural. Eng. 7 (2010), 046011. PMID: 20644246.
[30] S.M. Wellman, et al., Revealing spatial and temporal patterns of cell death, glial
proliferation, and blood-brain barrier dysfunction around implanted intracortical
neural interfaces, Front. Neurosci. 13 (2019) 493. PMID: 31191216.
[31] R. Biran, et al., Neuronal cell loss accompanies the brain tissue response to
chronically implanted silicon microelectrode arrays, Exp. Neurol. 195 (2005)
115–126. PMID: 16045910.
[32] G.C. McConnell, et al., Implanted neural electrodes cause chronic, local
inflammation that is correlated with local neurodegeneration, J. Neural. Eng. 6
(2009) (PMID).
[33] J.K. Hermann, et al., The role of toll-like receptor 2 and 4 innate immunity
pathways in intracortical microelectrode-induced neuroinflammation, Front.
Bioeng. Biotechnol. 6 (2018) 113. PMID: 30159311.
[34] J.K. Hermann, et al., Inhibition of the cluster of differentiation 14 innate
immunity pathway with IAXO-101 improves chronic microelectrode
performance, J. Neural. Eng. 15 (2018), 025002. PMID: 29219114.
[35] H.W. Bedell, et al., Differential expression of genes involved in the acute innate
immune response to intracortical microelectrodes, Acta Biomater. 102 (2020)
205–219. PMID: 31733330.
[36] G. Hajishengallis, J.D. Lambris, More than complementing Tolls: complementToll-like receptor synergy and crosstalk in innate immunity and inflammation,
Immunol. Rev. 274 (2016) 233–244. PMID: 27782328.
[37] J.G. Walsh, et al., Inflammasomes in the CNS, Nat. Rev. Neurosci. 15 (2014)
84–97. PMID: 24399084.
[38] L. Song, et al., NLRP3 inflammasome in neurological diseases, from functions to
therapies, Front. Cell. Neurosci. 11 (2017) 63. PMID: 28337127.
[39] S. Voet, et al., Inflammasomes in neuroinflammatory and neurodegenerative
diseases, EMBO Mol. Med. 11 (2019). PMID: 31015277.
[40] J.P. de Rivero Vaccari, et al., Therapeutics targeting the inflammasome after
central nervous system injury, Transl. Res. 167 (2016) 35–45. PMID: 26024799.
[41] J.P. de Rivero Vaccari, et al., Activation and regulation of cellular
inflammasomes: gaps in our knowledge for central nervous system injury,
J. Cerebr. Blood Flow Metabol. : Off. J. Int. Soc. Cerebral Blood Flow Metabol. 34
(2014) 369–375. PMID: 24398940.
[42] N. Kawana, et al., Reactive astrocytes and perivascular macrophages express
NLRP3 inflammasome in active demyelinating lesions of multiple sclerosis and
necrotic lesions of neuromyelitis optica and cerebral infarction, Clin.
Experimental Neuroimmunol. 4 (2013) 296–304 (PMID).
[43] S.W. Lee, et al., The role of microglial inflammasome activation in pyroptotic cell
death following penetrating traumatic brain injury, J. Neuroinflammation 16
(2019) 27. PMID: 30736791.
[44] J. Minkiewicz, et al., Human astrocytes express a novel NLRP2 inflammasome,
Glia 61 (2013) 1113–1121. PMID: 23625868.
[45] J.P. de Rivero Vaccari, et al., A molecular platform in neurons regulates
inflammation after spinal cord injury, J. Neurosci. 28 (2008) 3404–3414. PMID:
18367607.
[46] B. Cyr, et al., The role of non-canonical and canonical inflammasomes in
inflammaging, Front. Mol. Neurosci. 15 (2022), 774014. PMID: 35221912.
[47] N. Kerr, et al., Inflammasome proteins in Serum and serum-derived extracellular
vesicles as biomarkers of stroke, Front. Mol. Neurosci. 11 (2018) 309. PMID:
30233311.
[48] N. Kerr, et al., Inflammasome proteins as biomarkers of traumatic brain injury,
PLoS One 13 (2018), e0210128. PMID: 30596792.
[49] J. Perez-Barcena, et al., Levels of caspase-1 in cerebrospinal fluid of patients with
traumatic brain injury: correlation with intracranial pressure and outcome,
J. Neurosurg. 134 (2020) 1644–1649. PMID: 32357337.
[50] J. Perez-Barcena, et al., Serum caspase-1 as an independent prognostic factor in
traumatic brain injured patients, Neurocritical Care 36 (2022) 527–535. PMID:
34498205.
[51] S.E. Adamczak, et al., Pyroptotic neuronal cell death mediated by the AIM2
inflammasome, J. Cerebr. Blood Flow Metabol. : Off. J. Int. Soc. Cerebral Blood
Flow Metabol. 34 (2014) 621–629. PMID: 24398937.
[52] J.C. de Rivero Vaccari, et al., The inflammasome in times of COVID-19, Front.
Immunol. 11 (2020), 583373. PMID: 33149733.
[53] V. Govindarajan, et al., Role of inflammasomes in multiple sclerosis and their
potential as therapeutic targets, J. Neuroinflammation 17 (2020) 260. PMID:
32878648.
The project described was supported by Grant Number
UL1TR002736, Miami Clinical and Translational Science Institute, from
the National Center for Advancing Translational Sciences and the Na­
tional Institute on Minority Health and Health Disparities, an R01 grant
from the NIH/NINDS to RWK and JPdRV (R01NS113969-01) and an
RF1 grant from the NIH/NINDS/NIA (1RF1NS125578-01) to WDD and
JPdRV. Its contents are solely the responsibility of the authors and do
not necessarily represent the official views of the NIH. Fig. 1 was created
with Biorender. The authors would like to thank Dr. Florian Solzbacher
and Rohit Sharma at the University of Utah for providing the nonfunctional Utah microelectrode arrays tested in this study.
Appendix A. Supplementary data
Supplementary data to this article can be found online at https://doi.
org/10.1016/j.biomaterials.2023.122102.
References
[1] A.B. Schwartz, et al., Brain-controlled interfaces: movement restoration with
neural prosthetics, Neuron 52 (2006) 205–220. PMID: 17015237.
[2] J.L. Collinger, et al., Neuroprosthetic technology for individuals with spinal cord
injury, J Spinal Cord Med 36 (2013) 258–272. PMID: 23820142.
[3] M.A. Cervera, et al., Brain-computer interfaces for post-stroke motor
rehabilitation: a meta-analysis, Ann Clin Transl Neurol 5 (2018) 651–663. PMID:
29761128.
[4] N.C. Swann, et al., Adaptive deep brain stimulation for Parkinson’s disease using
motor cortex sensing, J. Neural. Eng. 15 (2018), 046006. PMID: 29741160.
[5] S. Debnath, et al., Long-term stability of neural signals from microwire arrays
implanted in common marmoset motor cortex and striatum, Biomed Phys Eng
Express 4 (2018). PMID: 31011432.
[6] A. Prasad, et al., Abiotic-biotic characterization of Pt/Ir microelectrode arrays in
chronic implants, Front. Neuroeng. 7 (2014) 2. PMID: 24550823.
[7] A. Prasad, et al., Comprehensive characterization and failure modes of tungsten
microwire arrays in chronic neural implants, J. Neural. Eng. 9 (2012), 056015.
PMID: 23010756.
[8] J.C. Barrese, et al., Scanning electron microscopy of chronically implanted
intracortical microelectrode arrays in non-human primates, J. Neural. Eng. 13
(2016), 026003. PMID: 26824680.
[9] J.C. Barrese, et al., Failure mode analysis of silicon-based intracortical
microelectrode arrays in non-human primates, J. Neural. Eng. 10 (2013),
066014. PMID: 24216311.
[10] A.B. Ajiboye, et al., Restoration of reaching and grasping movements through
brain-controlled muscle stimulation in a person with tetraplegia: a proof-ofconcept demonstration, Lancet 389 (2017) 1821–1830. PMID: WOS:
000400549300031.
[11] J.P. Donoghue, et al., Assistive technology and robotic control using motor cortex
ensemble-based neural interface systems in humans with tetraplegia, J. Physiol.
579 (2007) 603–611 (PMID).
[12] L.R. Hochberg, et al., Reach and grasp by people with tetraplegia using a neurally
controlled robotic arm, Nature 485 (2012) 372–375. PMID: 22596161.
[13] L.R. Hochberg, et al., Neuronal ensemble control of prosthetic devices by a
human with tetraplegia, Nature 442 (2006) 164–171. PMID: 16838014.
[14] J.D. Simeral, et al., Neural control of cursor trajectory and click by a human with
tetraplegia 1000 days after implant of an intracortical microelectrode array,
J. Neural. Eng. 8 (2011), 025027. PMID: 21436513.
[15] P.D. Ganzer, et al., Restoring the sense of Touch using a sensorimotor
demultiplexing neural interface, Cell 181 (2020) 763–773.e712. PMID:
32330415.
[16] C. Bennett, et al., The complement cascade at the Utah microelectrode-tissue
interface, Biomaterials 268 (2021), 120583. PMID: 33310540.
[17] C. Bennett, et al., Neuroinflammation, oxidative stress, and blood-brain barrier
(BBB) disruption in acute Utah electrode array implants and the effect of
deferoxamine as an iron chelator on acute foreign body response, Biomaterials
188 (2019) 144–159. PMID: 30343257.
[18] C. Bennett, et al., Blood brain barrier (BBB)-disruption in intracortical silicon
microelectrode implants, Biomaterials 164 (2018) 1–10. PMID: 29477707.
[19] N.F. Nolta, et al., BBB leakage, astrogliosis, and tissue loss correlate with silicon
microelectrode array recording performance, Biomaterials 53 (2015) 753–762.
PMID: 25890770.
[20] K. Woeppel, et al., Explant analysis of Utah electrode arrays implanted in human
cortex for brain-computer-interfaces, Front. Bioeng. Biotechnol. 9 (2021),
759711. PMID: 34950640.
[21] J.W. Fawcett, R.A. Asher, The glial scar and central nervous system repair, Brain
Res. Bull. 49 (1999) 377–391. PMID: 10483914.
[22] J.P. Frampton, et al., Effects of glial cells on electrode impedance recorded from
neuralprosthetic devices in vitro, Ann. Biomed. Eng. 38 (2010) 1031–1047.
PMID: 20336824.
12
M.E. Franklin et al.
Biomaterials 297 (2023) 122102
[54] K.A. Kigerl, et al., Pattern recognition receptors and central nervous system
repair, Exp. Neurol. 258 (2014) 5–16. PMID: 25017883.
[55] J.K. Hermann, J.R. Capadona, Understanding the role of innate immunity in the
response to intracortical microelectrodes, Crit. Rev. Biomed. Eng. 46 (2018)
341–367. PMID: 30806249.
[56] J.P. de Rivero Vaccari, et al., Therapeutic neutralization of the NLRP1
inflammasome reduces the innate immune response and improves histopathology
after traumatic brain injury, J. Cerebr. Blood Flow Metabol. : Off. J. Int. Soc.
Cerebral Blood Flow Metabol. 29 (2009) 1251–1261. PMID: 19401709.
[57] T.D. Kozai, et al., Effects of caspase-1 knockout on chronic neural recording
quality and longevity: insight into cellular and molecular mechanisms of the
reactive tissue response, Biomaterials 35 (2014) 9620–9634. PMID: 25176060.
[58] M.A. Lopez-Ramirez, et al., Role of caspases in cytokine-induced barrier
breakdown in human brain endothelial cells, J. Immunol. 189 (2012) 3130–3139.
PMID: 22896632.
[59] J. Yang, et al., Mechanism of gasdermin D recognition by inflammatory caspases
and their inhibition by a gasdermin D-derived peptide inhibitor, Proc. Natl. Acad.
Sci. U. S. A. 115 (2018) 6792–6797. PMID: 29891674.
[60] D. Zhang, et al., Gasdermin D serves as a key executioner of pyroptosis in
experimental cerebral ischemia and reperfusion model both in vivo and in vitro,
J. Neurosci. Res. 97 (2019) 645–660. PMID: 30600840.
[61] S. Kumar, Mechanisms mediating caspase activation in cell death, Cell Death
Differ. 6 (1999) 1060–1066. PMID: 10578174.
[62] D.P. Abulafia, et al., Inhibition of the inflammasome complex reduces the
inflammatory response after thromboembolic stroke in mice, J. Cerebr. Blood
Flow Metabol. : Off. J. Int. Soc. Cerebral Blood Flow Metabol. 29 (2009) 534–544.
PMID: 19066616.
[63] J.P. de Rivero Vaccari, et al., Exosome-mediated inflammasome signaling after
central nervous system injury, J. Neurochem. 136 (Suppl 1) (2016) 39–48. PMID:
25628216.
[64] S. Song, et al., Neuroinflammatory gene expression analysis reveals pathways of
interest as potential targets to improve the recording performance of intracortical
microelectrodes, Cells 11 (2022). PMID: 35954192.
[65] Julian, G.S., de Oliveira, R.W., Perry, J.C., Tufik, S. & Chagas, J.R. Validation of
housekeeping genes in the brains of rats submitted to chronic intermittent
hypoxia, a sleep apnea model. PLoS One 9, e109902 (2014), PMID: 25289636.
[66] K.J. Livak, T.D. Schmittgen, Analysis of relative gene expression data using realtime quantitative PCR and the 2(-Delta Delta C(T)) Method, Methods 25 (2001)
402–408. PMID: 11846609.
[67] A. Moniri, et al., Framework for DNA quantification and outlier detection using
multidimensional standard curves, Anal. Chem. 91 (2019) 7426–7434. PMID:
31056898.
[68] M.J. Burns, et al., Standardisation of data from real-time quantitative PCR
methods - evaluation of outliers and comparison of calibration curves, BMC
Biotechnol. 5 (2005) 31. PMID: 16336641.
[69] X. Rao, et al., An improvement of the 2^(-delta delta CT) method for quantitative
real-time polymerase chain reaction data analysis, Biostat Bioinforma Biomath 3
(2013) 71–85. PMID: 25558171.
[70] H.J. Motulsky, R.E. Brown, Detecting outliers when fitting data with nonlinear
regression - a new method based on robust nonlinear regression and the false
discovery rate, BMC Bioinf. 7 (2006) 123. PMID: 16526949.
[71] H.L. Desu, et al., IC100: a novel anti-ASC monoclonal antibody improves
functional outcomes in an animal model of multiple sclerosis,
J. Neuroinflammation 17 (2020) 143. PMID: 32366256.
[72] P.A. Amstad, et al., Detection of caspase activation in situ by fluorochromelabeled caspase inhibitors, 612, 614, passim, Biotechniques 31 (2001) 608–610.
PMID: 11570504.
[73] E. Bedner, et al., Activation of caspases measured in situ by binding of
fluorochrome-labeled inhibitors of caspases (FLICA): correlation with DNA
fragmentation, Exp. Cell Res. 259 (2000) 308–313. PMID: 10942603.
[74] E.A. Slee, et al., Serial killers: ordering caspase activation events in apoptosis, Cell
Death Differ. 6 (1999) 1067–1074. PMID: 10578175.
[75] J. Ghaye, et al., Image thresholding techniques for localization of sub-resolution
fluorescent biomarkers, Cytometry 83 (2013) 1001–1016. PMID: 24105983.
[76] Y.R. Van Eycke, et al., Image processing in digital pathology: an opportunity to
solve inter-batch variability of immunohistochemical staining, Sci. Rep. 7 (2017),
42964. PMID: 28220842.
[77] R. Waller, et al., Iba-1-/CD68+ microglia are a prominent feature of ageassociated deep subcortical white matter lesions, PLoS One 14 (2019), e0210888.
PMID: 30682074.
[78] C.A. Finney, et al., A scalable, fully automated approach for regional
quantification of immunohistochemical staining of astrocytes in the rat brain,
J. Neurosci. Methods 348 (2021), 108994. PMID: 33176173.
[79] K.O. Lopes, et al., Microglial dystrophy in the aged and Alzheimer’s disease brain
is associated with ferritin immunoreactivity, Glia 56 (2008) 1048–1060. PMID:
18442088.
[80] W.J. Streit, Microglial activation and neuroinflammation in Alzheimer’s disease:
a critical examination of recent history, Front. Aging Neurosci. 2 (2010) 22.
PMID: 20577641.
[81] W.J. Streit, Q.S. Xue, Life and death of microglia, J. Neuroimmune Pharmacol. :
Off. J. Soc. NeuroImmune Pharmacol. 4 (2009) 371–379. PMID: 19680817.
[82] J. Liu, et al., Post-stroke treatment with argon attenuated brain injury, reduced
brain inflammation and enhanced M2 microglia/macrophage polarization: a
randomized controlled animal study, Crit. Care 23 (2019) 198. PMID: 31159847.
[83] F. Moro, et al., Efficacy of acute administration of inhaled argon on traumatic
brain injury in mice, Br. J. Anaesth. 126 (2021) 256–264. PMID: 32977957.
[84] A.D. Degenhart, et al., Histological evaluation of a chronically-implanted
electrocorticographic electrode grid in a non-human primate, J. Neural. Eng. 13
(2016), 046019. PMID: 27351722.
[85] N.A. Hamilton, et al., Statistical and visual differentiation of subcellular imaging,
BMC Bioinf. 10 (2009) 94. PMID: 19302715.
[86] M.A. Garcia-Cabezas, et al., Distinction of neurons, glia and endothelial cells in
the cerebral cortex: an algorithm based on cytological features, Front. Neuroanat.
10 (2016) 107. PMID: 27847469.
[87] I.P. Karve, et al., The contribution of astrocytes and microglia to traumatic brain
injury, Br. J. Pharmacol. 173 (2016) 692–702. PMID: 25752446.
[88] T. Goldmann, et al., Love and death: microglia, NLRP3 and the Alzheimer’s brain,
Cell Res. 23 (2013) 595–596. PMID: 23399597.
[89] S.P. Savya, et al., In vivo spatiotemporal dynamics of astrocyte reactivity
following neural electrode implantation, Biomaterials 289 (2022), 121784.
PMID: 36103781.
[90] R. Biran, et al., The brain tissue response to implanted silicon microelectrode
arrays is increased when the device is tethered to the skull, J. Biomed. Mater.
Res., Part A 82 (2007) 169–178 (PMID).
[91] C.H. Thompson, et al., Spatiotemporal patterns of gene expression around
implanted silicon electrode arrays, J. Neural. Eng. 18 (2021). PMID: 33780909.
[92] M.A. Anderson, et al., Astrocyte scar formation aids central nervous system axon
regeneration, Nature 532 (2016) 195–200. PMID: 27027288.
[93] H. Guo, et al., Inflammasomes: mechanism of action, role in disease, and
therapeutics, Nat. Med. 21 (2015) 677–687. PMID: 26121197.
[94] Y. Sasaki, et al., Iba1 is an actin-cross-linking protein in macrophages/microglia,
Biochem. Biophys. Res. Commun. 286 (2001) 292–297. PMID: 11500035.
[95] S.W. Lee, et al., Microglial inflammasome activation in penetrating ballistic-like
brain injury, J. Neurotrauma 35 (2018) 1681–1693. PMID: 29439605.
[96] V. Parpura, et al., Glial cells in (patho)physiology, J. Neurochem. 121 (2012)
4–27. PMID: 22251135.
[97] J. Gehrmann, et al., Microglia: intrinsic immuneffector cell of the brain, Brain Res
Brain Res Rev 20 (1995) 269–287. PMID: 7550361.
[98] J.G. Walsh, et al., Rapid inflammasome activation in microglia contributes to
brain disease in HIV/AIDS, Retrovirology 11 (2014) 35. PMID: 24886384.
[99] M.K. Mamik, C. Power, Inflammasomes in neurological diseases: emerging
pathogenic and therapeutic concepts, Brain 140 (2017) 2273–2285. PMID:
29050380.
[100] D. Tang, et al., PAMPs and DAMPs: signal 0s that spur autophagy and immunity,
Immunol. Rev. 249 (2012) 158–175. PMID: 22889221.
[101] M. Schnaars, et al., Assessing beta-amyloid-induced NLRP3 inflammasome
activation in primary microglia, Methods Mol. Biol. 1040 (2013) 1–8. PMID:
23852592.
[102] L. Freeman, et al., NLR members NLRC4 and NLRP3 mediate sterile
inflammasome activation in microglia and astrocytes, J. Exp. Med. 214 (2017)
1351–1370. PMID: 28404595.
[103] J.P. de Rivero Vaccari, et al., P2X4 receptors influence inflammasome activation
after spinal cord injury, J. Neurosci. 32 (2012) 3058–3066. PMID: 22378878.
[104] N.A. Kerr, et al., Traumatic brain injury-induced acute lung injury: evidence for
activation and inhibition of a neural-respiratory-inflammasome Axis,
J. Neurotrauma 35 (2018) 2067–2076. PMID: 29648974.
[105] S.L. Chang, et al., NLRP12 inflammasome expression in the rat brain in response
to LPS during morphine tolerance, Brain Sci. 7 (2017). PMID: 28178176.
[106] I. Sastalla, et al., Transcriptional analysis of the three Nlrp1 paralogs in mice,
BMC Genom. 14 (2013) 188. PMID: 23506131.
[107] K.M. Cirelli, et al., Inflammasome sensor NLRP1 controls rat macrophage
susceptibility to Toxoplasma gondii, PLoS Pathog. 10 (2014), e1003927. PMID:
24626226.
[108] J.A. Duncan, S.W. Canna, The NLRC4 inflammasome, Immunol. Rev. 281 (2018)
115–123. PMID: 29247997.
[109] N.H. Mejias, et al., Contribution of the inflammasome to inflammaging,
J. Inflamm. 15 (2018) 23. PMID: 30473634.
[110] A. Denes, et al., AIM2 and NLRC4 inflammasomes contribute with ASC to acute
brain injury independently of NLRP3, Proc. Natl. Acad. Sci. U. S. A. 112 (2015)
4050–4055. PMID: 25775556.
[111] H. Matsuyama, et al., Chronic cerebral hypoperfusion activates AIM2 and NLRP3
inflammasome, Brain Res. 1736 (2020), 146779. PMID: 32171704.
[112] W.J. Rui, et al., Microglial AIM2 alleviates antiviral-related neuro-inflammation
in mouse models of Parkinson’s disease, Glia 70 (2022) 2409–2425. PMID:
35959803.
[113] W.E. Barclay, et al., The AIM2 inflammasome is activated in astrocytes during the
late phase of EAE, JCI Insight 7 (2022). PMID: 35451371.
[114] H. Kim, et al., AIM2 inflammasome contributes to brain injury and chronic poststroke cognitive impairment in mice, Brain Behav. Immun. 87 (2020) 765–776.
PMID: 32201254.
[115] S.Y. Xu, et al., AIM2 deletion enhances blood-brain barrier integrity in
experimental ischemic stroke, CNS Neurosci. Ther. 27 (2021) 1224–1237. PMID:
34156153.
[116] S.L. Lage, et al., Emerging concepts about NAIP/NLRC4 inflammasomes, Front.
Immunol. 5 (2014) 309. PMID: 25071770.
[117] M.S. Dick, et al., ASC filament formation serves as a signal amplification
mechanism for inflammasomes, Nat. Commun. 7 (2016), 11929. PMID:
27329339.
[118] A. Stutz, et al., ASC speck formation as a readout for inflammasome activation,
Methods Mol. Biol. 1040 (2013) 91–101. PMID: 23852599.
13
M.E. Franklin et al.
Biomaterials 297 (2023) 122102
[132] X. Liu, N. Quan, Microglia and CNS interleukin-1: beyond immunological
concepts, Front. Neurol. 9 (2018) 8. PMID: 29410649.
[133] G. Fenini, et al., Potential of IL-1, IL-18 and inflammasome inhibition for the
treatment of inflammatory skin diseases, Front. Pharmacol. 8 (2017) 278. PMID:
28588486.
[134] N. Kayagaki, et al., Non-canonical inflammasome activation targets caspase-11,
Nature 479 (2011) 117–121. PMID: 22002608.
[135] P. Gurung, et al., FADD and caspase-8 mediate priming and activation of the
canonical and noncanonical Nlrp3 inflammasomes, J. Immunol. 192 (2014)
1835–1846. PMID: 24453255.
[136] L. Bossaller, et al., Cutting edge: FAS (CD95) mediates noncanonical IL-1beta and
IL-18 maturation via caspase-8 in an RIP3-independent manner, J. Immunol. 189
(2012) 5508–5512. PMID: 23144495.
[137] N.H. Johnson, et al., Inflammasome Activation in Traumatic Brain Injury and
Alzheimer’s Disease, Transl Res, 2022. PMID: 36070840.
[138] S. Alboni, et al., Interleukin 18 in the CNS, J. Neuroinflammation 7 (2010) 9.
PMID: 20113500.
[139] S. Jander, et al., Interleukin-18 expression after focal ischemia of the rat brain:
association with the late-stage inflammatory response, J. Cerebr. Blood Flow
Metabol. : Off. J. Int. Soc. Cerebral Blood Flow Metabol. 22 (2002) 62–70. PMID:
11807395.
[140] S. Das, et al., Japanese Encephalitis Virus infection induces IL-18 and IL-1beta in
microglia and astrocytes: correlation with in vitro cytokine responsiveness of glial
cells and subsequent neuronal death, J. Neuroimmunol. 195 (2008) 60–72. PMID:
18374991.
[141] X. Ge, et al., The pathological role of NLRs and AIM2 inflammasome-mediated
pyroptosis in damaged blood-brain barrier after traumatic brain injury, Brain Res.
1697 (2018) 10–20. PMID: 29886252.
[119] J. Masumoto, et al., Pyrin N-terminal homology domain- and caspase recruitment
domain-dependent oligomerization of ASC, Biochem. Biophys. Res. Commun. 280
(2001) 652–655. PMID: 11162571.
[120] V. Sagulenko, et al., AIM2 and NLRP3 inflammasomes activate both apoptotic and
pyroptotic death pathways via ASC, Cell Death Differ. 20 (2013) 1149–1160.
PMID: 23645208.
[121] D.P. Sester, et al., A novel flow cytometric method to assess inflammasome
formation, J. Immunol. 194 (2015) 455–462. PMID: 25404358.
[122] U. Felderhoff-Mueser, et al., IL-18: a key player in neuroinflammation and
neurodegeneration? Trends Neurosci. 28 (2005) 487–493. PMID: 16023742.
[123] X.O. Scott, et al., The inflammasome adaptor protein ASC in mild cognitive
impairment and alzheimer’s disease, Int. J. Mol. Sci. 21 (2020). PMID: 32630059.
[124] S.H. Chen, et al., Netosis and inflammasomes in large vessel occlusion thrombi,
Front. Pharmacol. 11 (2020), 607287. PMID: 33569001.
[125] S.J. Hewett, et al., Interleukin-1beta in central nervous system injury and repair,
Eur J Neurodegener Dis 1 (2012) 195–211. PMID: 26082912.
[126] C.A. Dinarello, et al., Interleukin-18 and IL-18 binding protein, Front. Immunol. 4
(2013) 289. PMID: 24115947.
[127] T. Menge, et al., Induction of the proinflammatory cytokine interleukin-18 by
axonal injury, J. Neurosci. Res. 65 (2001) 332–339. PMID: 11494369.
[128] B. Conti, et al., Cultures of astrocytes and microglia express interleukin 18, Brain
Res Mol Brain Res 67 (1999) 46–52. PMID: 10101231.
[129] R.D. Wheeler, et al., Interleukin-18 induces expression and release of cytokines
from murine glial cells: interactions with interleukin-1 beta, J. Neurochem. 85
(2003) 1412–1420. PMID: 12787061.
[130] G. Lopez-Castejon, D. Brough, Understanding the Mechanism of IL-1beta
Secretion, vol. 22, Cytokine Growth Factor Rev, 2011, pp. 189–195. PMID:
22019906.
[131] B.R. Barker, et al., Cross-regulation between the IL-1beta/IL-18 processing
inflammasome and other inflammatory cytokines, Curr. Opin. Immunol. 23
(2011) 591–597. PMID: 21839623.
14