Skip to content

Advertisement

  • Research
  • Open Access

Complement components are upregulated and correlate with disease progression in the TDP-43Q331K mouse model of amyotrophic lateral sclerosis

Journal of Neuroinflammation201815:171

https://doi.org/10.1186/s12974-018-1217-2

©  The Author(s). 2018

  • Received: 5 March 2018
  • Accepted: 24 May 2018
  • Published:

Abstract

Background

Components of the innate immune complement system have been implicated in the pathogenesis of amyotrophic lateral sclerosis (ALS) specifically using hSOD1 transgenic animals; however, a comprehensive examination of complement expression in other transgenic ALS models has not been performed. This study therefore aimed to determine the expression of several key complement components and regulators in the lumbar spinal cord and tibialis anterior muscle of TDP-43Q331K mice during different disease ages.

Methods

Non-transgenic, TDP-43WT and TDP-43Q331K mice were examined at three different ages of disease progression. Expression of complement components and their regulators were examined using real-time quantitative PCR and enzyme-linked immunosorbent assay. Localisation of terminal complement component receptor C5aR1 within the lumbar spinal cord was also investigated using immunohistochemistry.

Results

Altered levels of several major complement factors, including C5a, in the spinal cord and tibialis anterior muscle of TDP-43Q331K mice were observed as disease progressed, suggesting overall increased complement activation in TDP-43Q331K mice. C5aR1 increased during disease progression, with immuno-localisation demonstrating expression on motor neurons and expression on microglia surrounding the regions of motor neuron death. There was a strong negative linear relationship between spinal cord C1qB, C3 and C5aR1 mRNA levels with hind-limb grip strength.

Conclusions

These results indicate that similar to SOD1 transgenic animals, local complement activation and increased expression of C5aR1 may contribute to motor neuron death and neuromuscular junction denervation in the TDP-43Q331K mouse ALS model. This further validates C5aR1 as a potential therapeutic target for ALS.

Background

Amyotrophic lateral sclerosis (ALS), also known as motor neuron disease, is a devastatingly fatal neurodegenerative disorder for which there are few effective treatments. ALS is characterised by loss of cortical spinal neurons of the motor cortex and alpha motor neurons within the brainstem and spinal cord, which results in skeletal muscle atrophy and progressive paralysis, eventually leading to death within 2 to 5 years of diagnosis. In the vast majority of ALS patients (~ 90%), the disease develops sporadically; however, in a minority of cases (~ 10%), the disease has a familial component, and it is due to specific genetic mutations. Some of the genes that have been implicated in ALS include C9orf72, VCP, FUS, SOD1 and TARDBP (TDP-43) [1–5]. Despite these differing aetiologies, sporadic and familial ALS patients are clinically and pathologically indistinguishable, suggesting that regardless of whether an ALS patient carries a known ALS mutation, or is sporadic, the underlying mechanism of motor neuron dysfunction is similar [6]. Numerous mechanisms have been proposed to contribute to ALS pathophysiology, including neuroinflammation. A key mediator of neuroinflammation is the chronic activation of the complement system, proposed to drive ALS disease progression [7].

Multiple clinical and experimental studies have shown compelling evidence that complement activation is involved in the pathogenesis of ALS, whereby components of all the complement pathways are upregulated in the serum, cerebrospinal fluid, skeletal muscles and neurological tissue (spinal cord and motor cortex) of ALS patients, as well as in transgenic SOD1 animal models of ALS [8]. Chronic complement activation is proposed to drive ALS disease progression through the actions of the pro-inflammatory complement peptide, C5a, signalling through its main receptor C5aR1 [9, 10]. This pathogenic role of C5a-C5aR1 is proposed to drive disease progression through inducing glial chemotaxis, activation of local immune cells and infiltration of macrophages into skeletal muscles, thereby inducing an overall increase in inflammation/neuroinflammation and thus neurodegeneration [9–12]. However, the pathogenic role of C5a-C5aR1 signalling in ALS has primarily been shown in transgenic SOD1 rodent models of ALS [9, 12]. Hence, it is unknown whether C5a-C5aR1 pathogenic signalling is specific to ALS cases not characterised by SOD1 pathology. Thus, as majority of ALS patients show TDP-43 pathology (~ 95%), the current study aimed to investigate complement in a recently developed TDP-43Q331K mouse model of ALS [13].

We examined the expression of major complement factors and of C5a and its receptor C5aR1, within the lumbar spinal cord and tibialis anterior (TA) leg muscle at three different ages during disease progression in TDP-43Q331K mice, in order to provide a comprehensive overview of the potential involvement of complement in an alternative mouse model of ALS. Our findings demonstrate that a global dysregulation of complement system is involved in this TDP-43 familial mouse model of ALS, suggesting that complement/C5aR1 could be a potential therapeutic target in most forms of ALS.

Methods

Animals

Transgenic TDP-43WT (Line 96) and TDP-43Q331K (Line 103) mice were obtained from the Jackson Laboratory (Bar Harbor, ME, USA) and were bred on a C57BL/6J background to produce TDP-43WT, TDP-43Q331K and respective non-transgenic (NTg) control mice. TDP-43WT transgenic mice express a myc-tagged human non-mutated version of the TDP-43 cDNA sequence and TDP-43Q331K mice express a myc-tagged, human TDP-43 cDNA modified to have an ALS-linked glutamine to lysine residue mutation at position 331, under the direction of the mouse prion protein promoter. The prion protein promoter ensures that the transgene expression is directed primarily to the central nervous system—the brain and spinal cord—and is very low in other peripheral tissues [13]. Female NTg, TDP-43WT and TDP-43Q331K mice at 3, 10 months and 16 months based on their motor deficits were used in this study [14]. TDP-43WT mice have moderate overexpression levels in total TDP-43, with a 1.5-fold increase in total TDP-43 expression and similar levels of human wild-type TDP-43 levels compared with endogenous TDP-43 in NTg mice. TDP-43Q331K mice also have moderate overexpression levels in total TDP-43, with a 2.5-fold increase in total TDP-43 expression and 1.5-fold greater expression of human Q331K TDP-43 levels compared with endogenous TDP-43 in NTg mice [13].

Hind-limb grip strength test

The hind-limb grip strength of NTg, TDP-43WT and TDP-43Q331K female mice (n = 15 per genotype) was measured by using a grip strength meter (IMADA, Toyohashi, Japan) at 3, 10 and 16 months of age at the same time (14:00 h) of the day as previously described by us [9, 15, 16]. In brief, mice were held by their tails and lowered until they grasped the T-bar connected to the digital force transducer with their hind-limbs. The tail was lowered until the body was horizontal, and the mouse was pulled away from the T-bar with a smooth steady pull until both hind-limbs released the T-bar. The strength of the grip was measured in newtons. Each mouse was given 10 attempts, and the maximum strength was recorded [9].

Tissue preparation for microglia/astrocyte quantification and immunohistochemistry

Female NTg, TDP-43WT and TDP-43Q331K mice (n = 4 per genotype) were euthanized by intraperitoneal injection of zolazapam (50 mg/kg; Zoletil, Lyppard) and xylazine (10 mg/kg; Xylazil, Lyppard). Mice were then fixed by transcardiac perfusion with 2% sodium nitrite in 0.1 M phosphate buffer (pH 7.4; Sigma-Aldrich, St Louis, MO, USA) followed by 4% paraformaldehyde in 0.1 M phosphate buffer (4% PFA-PB; pH 7.4; Sigma-Aldrich, St Louis, MO, USA) at the previously mentioned ages. Lumbar spinal cords were collected and placed into 4% PFA-PB for 2 h at 4 °C. Following this incubation, the spinal cords were washed 3 × 5 min in phosphate-buffered saline (PBS; pH 7.4), followed by submersion in sucrose solution at 15% then 30% in PBS (pH 7.4). Lumbar spinal cords were then embedded in optimal cutting temperature compound (Sakura, Finetek, Torrance, CA, USA) then snap frozen in liquid nitrogen. Lumbar spinal cords were sectioned into 16-μm-thick transverse and coronal sections and dry mounted onto Superfrost Plus slides (Menzel-Glaser, Braunschweig, Germany) for estimation of astrocytes, microglia and immunohistochemistry as detailed below.

Estimation of astrocytes and microglia

For estimation of astrocytes and microglia within the lumbar spinal cord, sections were rehydrated in PBS (pH 7.4) then blocked in PBS containing 3% bovine serum albumin (BSA) for 1 h at room temperature. Sections were incubated overnight at 4 °C with the astrocyte (mouse anti-GFAP; 1:1000, BD Biosciences, San Diego, CA, USA) and microglia (rat anti-CD11b; 1:500, Abcam, Cambridge, MA, USA) markers. Sections were washed with PBS for 3 × 10 min prior to incubation overnight at 4 °C with the Alexa secondary cocktail: Alexa Fluor 555 dye-conjugated goat anti-rat (1:1000, Invitrogen, Eugene, OR, USA) and Alexa Fluor 488 dye-conjugated goat anti-mouse (1:600, Invitrogen, Eugene, OR, USA) antibody. All primary and secondary antibodies were diluted in PBS (pH 7.4) containing 1% BSA. Sections were then washed for 3 × 5 min in PBS, then mounted with Prolong Gold Anti-Fade medium containing 4, 6-diamidino-2-phenylindole (DAPI; Invitrogen, Eugene, OR, USA). Quantification of GFAP and CD11b was performed on ~ 11 to 14 lumbar spinal cord sections spaced 320 μm apart and expressed as the percentage immunoreactive area per section [12]. Quantification was within the second lumbar dorsal root ganglia (L2) to the fifth lumbar dorsal root ganglia (L5), selected with the aid of the mouse spinal cord atlas [17]. Staining procedures and image exposures were all standardised between genotypes and between sections. The mouse genotype was not made available to the researchers until the completion of the study.

Quantification of activated microglia numbers

The cell body of microglia was labelled with the nuclear marker, DAPI. As microglia are known to display morphological changes when they become activated, such as an increase in cell body size, thickening of proximal processes and a decrease in the ramification of distal branches [18], activated microglia were defined by the presence of one DAPI stain, an amoeboid cell body and proximal processes length ≤ 1–2 μm [19]. The total number of activated microglia was determined by the average of 11–14 sections, with the overall average multiplied by the number of sections within L2–L5. The mouse genotype was not made available to the researchers until quantification was completed.

Immunohistochemistry

Fluorescence double-labelling immunohistochemistry was performed to localise C1q and C5aR1 expression with specific cell-type markers for astrocytes, microglia and motor neurons as previously described [15]. Briefly, sections were rehydrated in PBS (pH 7.4) for 10 min, then blocked in PBS containing 3% BSA or 3% donkey serum (DS) for 1 h at room temperature. Sections were incubated overnight at 4 °C with combination of primary antibodies outlined in Table 1. All primary antibodies were diluted in PBS (pH 7.4) containing 1% BSA or 1% DS. Sections were washed 3 × 10 min with PBS prior to incubation with an appropriate Alexa-conjugated secondary cocktail: Alexa 555 goat anti-rat, Alexa 594 donkey anti-rat, Alexa 488 goat anti-mouse, Alexa 488 goat anti-rabbit and Alexa 488 donkey anti-goat (Invitrogen, Eugene, OR, USA). All secondary antibodies were diluted in PBS (pH 7.4) containing 1% BSA or 1% DS (1:1,1000 for Alexa 555, 1:500 for Alexa 594 and 1:600 for Alexa 488). Following 3 × 5 min washes in PBS, all sections were mounted with Prolong Gold Anti-Fade medium containing DAPI (Invitrogen, Eugene, OR, USA). Sections with no primary antibodies were used as negative controls for all immunohistochemistry experiments to give a measure of non-specific background staining.
Table 1

Summary of antibodies used for immunohistochemistry

Antibody

Manufacturer

Dilution

In combination with one of the following antibodies

Rat anti-mouse C5aR1

Bio-Rad

1:250

GFAP, Iba-1 or ChAT

Rat anti-mouse C1q

Hycult Biotechnology

1:1000

GFAP, Iba-1 or ChAT

Goat anti-mouse ChAT

Merck

1:100

C1q or C5aR1

Rabbit anti-mouse Iba-1

Wako

1:400

C1q or C5aR1

Mouse anti-rat GFAP

BD Biosciences

1:1000

C1q or C5aR1

Quantification of immunofluorescence for C1q was performed on ~ 25 to 35 lumbar spinal cord sections (per animal; n = 4) spaced 160 μm apart and expressed as the percentage immunoreactive area per section. Staining procedures and image exposures were all standardised between genotype and between sections [15].

Real-time quantitative PCR

Total RNA was isolated from the spinal cord and TA muscle of NTg, TDP-43WT and TDP-43Q331K mice using RNeasy Lipid Tissue extraction kit according to manufacturer’s instructions (QIAGEN, CA, USA). Total RNA was purified from genomic DNA contamination using Turbo DNAse treatment (Ambion, NY, USA), then converted to cDNA using AffinityScript cDNA synthesis kit according to manufacturer’s instructions (Agilent Technologies, CA, USA). Commercially available gene-specific Taqman probes for complement component 1, q subcomponent, beta polypeptide (C1qB; Mm01179619_m1), complement component 4 (C4; Mm00437893_g1), complement factor B (Cfb; Mm00433909_m1), complement component 3 (C3; Mm01232779_m1), CD55 antigen (Cd55; Mm00438377_m1), CD59a antigen (Cd59a; Mm00483149_m1) and complement component 5a receptor 1 (C5ar1; Mm00500292_s1) were used to amplify target gene of interest (Applied Biosystems, MA, USA). Relative target gene expression to geometric mean of reference genes glyceraldehyde-3-phosphate dehydrogenase (Gapdh; Mm99999915_g1), beta actin (Actb; Mm02619580_g1) and hypoxanthine guanine phosphoribosyl transferase (Hprt; Mm03024075_m1) was determined using this formula: 2-∆CT where ∆CT = (Ct (Target gene) − Ct (Gapdh, Actb and Hprt)), as per our previous studies [15, 16]. Final measures are presented as relative levels of gene expression in TDP-43WT and TDP-43Q331K mice compared with expression in NTg controls. Probe sets were tested over a serial cDNA concentration for amplification efficiency. No reverse transcription and water as no template control was used as negative controls. All samples were run in triplicate and were tested in three separate experiments.

Enzyme-linked immunosorbent assay

Ninety-six-well plates (Greiner Bio-One, Frickenhausen, Germany) were pre-coated with rat anti-mouse C5a capture antibody (4 μg/mL; R&D Systems, Minneapolis, MN, USA) diluted in PBS (pH 7.4) overnight at room temperature in a sealed humidified container. Following the plate being blocked for 1 h at room temperature with reagent diluent (1% BSA in PBS), C5a standard, spinal cord and TA muscle homogenates were incubated for 2 h at room temperature. The plates were subsequently incubated with biotinylated goat anti-mouse C5a detection antibody (0.2 μg/mL, R&D Systems, Minneapolis, MN, USA) for 1 h at room temperature, and then incubated with Streptavidin-HRP conjugate for 20 min at room temperature. Tetramethylbenzidine (Sigma-Aldrich, Saint Louis, MO, USA) substrate was used as the chromogen, and the plate was read at 450 nm. Levels of C5a in spinal cord and TA muscle samples were adjusted to micrograms per protein and expressed as nanograms of C5a per microgram of protein.

Statistical analysis

All statistical analyses were performed using GraphPad Prism 7.0 (GraphPad Software Inc., San Diego, CA, USA). For the results from GFAP and CD11b quantification, quantitative real-time PCR and enzyme-linked immunosorbent assay, statistical differences between NTg, TDP-43WT and TDP-43Q331K mice were determined using one-way ANOVA with Tukey’s post hoc test for each age group. To assess the linear association between complement mRNA transcript levels and hind-limb grip strength of NTg, TDP-43WT and TDP-43Q331K mice, Pearson’s correlation was used. All data was presented as mean ± standard error of mean and differences considered significant when P < 0.05.

Results

Motor deficits in TDP-43Q331K mice correlate with increases in astrocytes and microglia during disease progression

To monitor the decline in motor performance during disease onset and progression in TDP-43Q331K mice, we performed hind-limb grip strength tests in animals. At 3 months, there was no difference in hind-limb grip strength between NTg, TDP-43WT and TDP-43Q331K mice (Fig. 1a). However, by 10 months, we observed a significant reduction in hind-limb grip strength in TDP-43Q331K mice when compared with NTg and TDP-43WT mice (~ 30% reduction, n = 15, ****P < 0.0001, ++++P < 0.0001; Fig. 1a). Furthermore, at 16 months, there was a progressive decline in hind-limb grip strength in TDP-43Q331K mice (~ 45% reduction, n = 15, ****P < 0.0001, ++++P < 0.0001; Fig. 1a) when compared with NTg and TDP-43WT mice. Importantly, we found that the decline in hind-limb grip strength in TDP-43Q331K mice closely correlated with the increase in immunoreactive area of astrocytes using immunofluorescence staining in the lumbar spinal cords at 10 months (~ 200% increase, n = 4, **P < 0.01; Fig. 1b, c) and 16 months (~ 380% increase, n = 4, ****P < 0.0001; Fig. 1b, c). Similarly, we also found that the decline in hind-limb grip strength in TDP-43Q331K mice closely correlated with the increase in immunoreactive area of microglia and number of activated microglia in the lumbar spinal cord at 10 months (140~230% increase, n = 4, **P < 0.01, ***P < 0.001, ****P < 0.0001; Fig. 1d–f) and 16 months (130~280% increase, n = 4, *P < 0.05, **P < 0.01, ***P < 0.001, ****P < 0.0001; Fig. 1d–f). Taken together, these data reveal an age-related decline in hind-limb grip strength of TDP-43Q331K mice associated with increased glial activation in the lumbar spinal cord.
Fig. 1
Fig. 1

Decline in hind-limb grip strength during ALS progression correlates with increase in astrocytes and microglia in the lumbar spinal cord of TDP-43Q331K mice. a Hind-limb grip strength (N) from NTg, TDP-43WT and TDP-43Q331K mice at 3, 10 and 16 months of age. At 3 months, no significant difference in force is present between genotypes. By 10 months, TDP-43Q331K mice display a significant decrease in hind-limb grip strength compared with both NTg and TDP-43WT mice (~ 30% reduction; NTg vs TDP-43Q331K ****P < 0.0001 and TDP-43WT vs TDP-43Q331K ++++P < 0.0001; n = 15). A progressive decline in hind-limb grip strength is present at 16 months, with TDP43Q331K mice showing significantly lower hind-limb grip strength compared with NTg and TDP-43WT mice (~ 45% reduction; NTg vs TDP-43Q331K ****P < 0.0001 and TDP-43WT vs TDP-43Q331K ++++P < 0.0001; n = 15). b Representative images of the GFAP-positive astrocytes in the lumbar spinal cord of NTg, TDP-43WT and TDP-43Q331K animals at 16 months. Dashed line shows the outline of the ventral horn with higher magnification of the white square. Scale bars = 100 μm. c Increased astrocyte expression in TDP-43Q331K mice (blue bars) compared with NTg (orange bars) and TDP-43WT mice (green bars) at ages 10 and 16 months (**P < 0.01, ****P < 0.0001; n = 4). d Representative images of CD11b-positive microglia in the lumbar spinal cord of NTg, TDP-43WT and TDP-43Q331K animals at 16 months. Dashed line shows the outline of the ventral horn with higher magnification of the white square. Scale bar = 100 μm. e Increased microglia expression in TDP-43Q331K mice (blue bars) compared with NTg (orange bars) and TDP-43WT mice (green bars) at ages 10 and 16 months (*P < 0.05, **P < 0.01; n = 4). f Increased number of activated microglia (amoeboid) in TDP-43Q331K mice (blue bars) compared with NTg (orange bars) and TDP-43WT (green bars) mice at 10 and 16 months (***P < 0.001, ****P < 0.0001; n = 4). Data are presented as mean ± SEM; one-way ANOVA with Tukey’s post hoc test for each age

Components of the classical/lectin pathways of complement are upregulated along with decreased expression levels of complement regulator CD55 in TDP-43Q331K mice

The complement system is part of the innate immune system that can contribute to neuroinflammation in many neurodegenerative diseases, including ALS [8]. Previous studies, including our own, have identified major complement components are upregulated in the lumbar spinal cord of hSOD1G93A mice [15]. However, there is no comprehensive overview of complement system in different animal models of ALS other than hSOD1 transgenic mice [8]. Therefore, we measured the mRNA levels of key components of the classical/lectin pathway (C1qB and C4), alternative pathway (fB), the central component to all pathways (C3) and the complement regulators (CD55 and CD59a) in the lumbar spinal cord of NTg, TDP-43WT and TDP-43Q331K mice using quantitative real-time PCR during disease progression of ALS (3, 10 and 16 months).

Quantitative real-time PCR analyses showed significant increases of the C1qB transcript in TDP-43Q331K mice by 1.3-fold at 10 months of age when compared to NTg and TDP-43WT mice; at 16 months of age, the increase was 1.5-fold and 1.4-fold when compared to NTg and TDP-43WT mice, respectively (blue bar compared to orange and green bars respectively in Fig. 2a; n = 5, ***P < 0.001, ****P < 0.0001). C4 transcript was also increased by 2.4-fold and 3.3-fold at 10 months of age and 2.0-fold and 2.7-fold at 16 months of age when compared to NTg and TDP-43WT mice, respectively (blue bar compared to orange and green bars respectively in Fig. 2b; n = 5, ***P < 0.001, ****P < 0.0001). By contrast, fB did not show any significant changes at 10 and 16 months of age in TDP-43Q331K mice when compared to NTg and TDP-43WT mice (blue bar compared to orange and green bars respectively in Fig. 2c; n = 5, P > 0.05). The central component of complement, C3, was also increased in the lumbar spinal cord of TDP-43Q331K mice, with a 1.6-fold and 1.4-fold increase at 10 months of age when compared to NTg and TDP-43WT mice, respectively; at 16 months of age, the increase was 1.7-fold when compared to both genotypes (blue bar compared to orange and green bars respectively in Fig. 2d; n = 5, ***P < 0.001, ****P < 0.0001).
Fig. 2
Fig. 2

Dysregulation of complement components in the lumbar spinal cord of TDP-43Q331K mice at three different ages of disease progression. a–f The mRNA expression profiles of the following complement components: C1qB (a, classical pathway), C4 (b, classical/lectin pathway), fB (c, alternative pathway), C3 (d, central component), CD55 (e, regulator) and CD59a (f, regulator) in the lumbar spinal cord of TDP-43Q331K mice (blue bars) relative to non-transgenic (NTg, orange bars) and TDP-43WT (green bars) mice during 3, 10 and 16 months of age. Data are expressed as means ± SEM (n = 5 mice/group, *P < 0.05, **P < 0.01, ***P < 0.001, ****P < 0.0001, one-way ANOVA with Tukey’s post hoc test)

The negative regulators of the complement system, CD55 and CD59a, were also investigated due to their importance in maintaining homeostasis and keeping the complement system in its proper physiological state in response to altered physiological conditions (i.e. infection and/or neurodegeneration). CD55 mRNA expression in TDP-43Q331K mice was decreased at 10 months of age by 0.3-fold and 0.4-fold, and by 0.4-fold and 0.5-fold at 16 months of age, when compared with NTg and TDP-43WT mice (blue bar compared to orange and green bars respectively in Fig. 2e; n = 5, *P < 0.05, **P < 0.01, ****P < 0.0001). By contrast, CD59a mRNA expression did not significantly alter at 10 and 16 months of age in TDP-43Q331K mice when compared to NTg and TDP-43WT mice (blue bar compared to orange and green bars respectively in Fig. 2f; n = 5, P > 0.05). These results suggest widespread complement perturbation occurs in the lumbar spinal cord of TDP-43Q331K mice, which may contribute to glial activation and neuroinflammation, and ultimately disease progression in this model.

Upregulation of C1q at 16 months of age was also confirmed using immunofluorescence, where there was marked increase in TDP-43Q331K mice compared with NTg and TDP-43WT mice (blue bar compared to orange and green bars in Fig. 3a; n = 4, *P < 0.05). We also observed that the marked increase of C1q in TDP-43Q331K mice was localised to motor neurons and microglia (white arrows in Fig. 3d, g), compared with NTg and TDP-43WT mice where little to no C1q was observed (Fig. 3b, c, e, f). We did not observe C1q on astrocytes in either NTg, TDP-43WT or TDP-43Q331K mice (Fig. 3h–j).
Fig. 3
Fig. 3

Expression and localisation of C1q in the spinal cord of non-transgenic, TDP-43WT and TDP-43Q331K mice at 16 months of age. a The immunoreactive area of C1q in the spinal cord of non-transgenic (NTg, orange bars), TDP-43WT (green bars) and TDP-43Q331K (blue bars) at 16 months. Data are expressed as means ± SEM (n = 4 mice/group, *P < 0.05, one-way ANOVA with Tukey’s post hoc test). b–j Double immunolabelling of C1q (red) with cellular markers (green) for motor neurons (ChAT; b–d), microglia (Iba-1; e–g) and astrocyte (GFAP; h–j) in the ventral lumbar spinal cord of NTg, TDP-43WT and TDP-43Q331K mice at 16 months of age. C1q was co-localised with ChAT-positive motor neurons (d, white arrow) and microglia (g, white arrow) in TDP-43Q331K mice. Note that white arrows indicate red and green fluorescent signal merge to orange. In TDP-43Q331K mice, immunolabelling of C1q was also evident on other cell types, indicated by lack of co-localisation with anti-ChAT, anti-Iba-1 and anti-GFAP (d, g, j). Scale bars for all panels = 20 μm

The terminal complement pathway C5a receptor, C5aR1, is upregulated and expressed on motor neurons and microglia in the lumbar spinal cord of TDP-43Q331K mice

Previous studies have shown increases in C5aR1 expression in the central nervous system of multiple rodent models of ALS, with many studies suggesting a pathogenic role for C5aR1 in the disease progression of ALS in hSOD1G93A mice [9, 10, 12, 20]. C5a, the ligand for C5aR1, is an activation fragment of the terminal complement cascade that is rapidly generated following complement cascade initiation [21]. We therefore examined the protein levels of C5a in the spinal cord of TDP-43Q331K mice using enzyme-linked immunosorbent assay as a biomarker for terminal complement activation. Interestingly, the results showed no change in C5a protein levels in TDP-43Q331K mice at 16 months of age when compared to NTg and TDP-43WT mice (blue bar compared to orange and green bars in Fig. 4a; n = 5, P > 0.05). In contrast to protein levels of C5a, C5aR1 mRNA expression was significantly increased by 1.4-fold and 1.2-fold at 10 months of age and by 1.8-fold and 1.6-fold at 16 months of age, when compared to NTg and TDP-43WT mice (blue bar compared to orange and green bars in Fig. 4b; n = 5, *P < 0.05, ***P < 0.001, ****P < 0.0001).
Fig. 4
Fig. 4

Expression of C5a and C5aR1 in the spinal cord of TDP-43Q331K mice at three different ages of disease progression. a The protein expression of C5a in the spinal cord of non-transgenic (NTg, orange bars), TDP-43WT (green bars) and TDP-43Q331K (blue bars) at 16 months. b mRNA expression of C5aR1 in the spinal cord of TDP-43Q331K mice relative to age-matched NTg and TDP-43WT mice at 3, 10 and 16 months of age. Data are expressed as means ± SEM (n = 5 mice/group, *P < 0.05, ***P < 0.001, ****P < 0.0001, one-way ANOVA with Tukey’s post hoc test for each age). c–k Double immunolabelling of C5aR1 (red) with cellular markers (green) for motor neurons (ChAT; c–e), microglia (Iba-1; f–h) and astrocyte (GFAP; i–k) in the ventral lumbar spinal cord of NTg, TDP-43WT and TDP-43Q331K mice at 16 months of age. C5aR1 was co-localised with ChAT-positive motor neurons (c–e, white arrows) and microglia (f–h, white arrows) in NTg, TDP-43WT and TDP-43Q331K mice. Note that white arrows indicate red and green fluorescent signal merge to orange. Scale bars for all panels = 20 μm

We next aimed to determine the cellular localisation of C5aR1 that could explain the increased expression seen in TDP-43Q331K mice. To achieve this, we performed immunolabelling for C5aR1 on the lumbar spinal cord from NTg, TDP-43WT and TDP-43Q331K mice. These sections were immunostained for C5aR1 with specific cellular markers to identify motor neurons (anti-ChAT), astrocytes (anti-GFAP) and microglia (anti-Iba-1). C5aR1 localised to ChAT-positive motor neurons and Iba-1-positive microglia in NTg, TDP-43WT and TDP-43Q331K mice at 16 months of age (white arrows in Fig. 4c–e for motor neurons and Fig. 4f–h for microglia), whereas it was not observed in GFAP-positive astrocytes (Fig. 4i–k). Together, the results above indicate that C5a-C5aR1 signalling could play a role in facilitating microglia activation and phagocytosis ultimately leading to motor neuron death in these animals.

Dysregulation of complement in the tibialis anterior muscle of TDP-43Q331K mice

In addition to showing altered levels of complement components in the spinal cord of TDP-43Q331K mice, we also investigated the level of major complement components in the TA muscle of TDP-43Q331K mice, as it has been shown previously in hSOD1G93A mice that complement is upregulated in this muscle. To investigate this, we measured the mRNA levels of key components of the complement pathways, which include the classical/lectin (C1qB and C4), alternative (fB) and terminal pathways (C5a and C5aR1), as well as the major complement regulators (CD55 and CD59a) using quantitative real-time PCR and enzyme-linked immunosorbent assay for C5a in TA muscle of TDP-43Q331K mice during disease progression of ALS (3, 10 and 16 months).

C1qB and C4 transcripts were significantly increased by 1.6-fold and 1.9-fold when compared to NTg mice and by 1.5-fold when compared to TDP-43WT mice at 10 months of age, respectively (blue bar compared to orange and green bars in Fig. 5a, b; n = 5, *P < 0.05, **P < 0.01, ***P < 0.001). C1qB and C4 transcripts were also increased by 1.7-fold and 1.5-fold when compared to NTg mice and by 1.7-fold and 1.9-fold when compared to TDP-43WT mice at 16 months of age, respectively (blue bar compared to orange and green bars in Fig. 5a, b; n = 5, *P < 0.05, **P < 0.01, ***P < 0.001). In addition to C1qB and C4, fB also displayed a marked increase in mRNA levels by 1.4-fold and 1.3-fold at 10 months of age when compared to NTg and TDP-43WT mice, whereas 1.5-fold increase at 16 months of age when compared to TDP-43WT mice (blue bar compared to orange and green bars in Fig. 5c; n = 5, *P < 0.05, **P < 0.01). By contrast, C3 was surprisingly decreased by 0.3-fold in TA muscle of TDP-43Q331K mice when compared to NTg and TDP-43WT mice at 10 months of age (blue bar compared to orange and green bars in Fig. 5d; n = 5, *P < 0.05, **P < 0.01). The regulator, CD55, was increased by 1.3-fold at 10 months of age in TDP-43Q331K mice when compared to NTg mice (blue bar compared to orange bar in Fig. 5e; n = 5, *P < 0.05), while there was no significant change in CD59a in TDP-43Q331K mice when compared to NTg and TDP-43WT controls (Fig. 5f; n = 5, P > 0.05). Lastly, the terminal complement pathway component, C5a, and its receptor C5aR1 were investigated. The results showed significant increases in C5a protein at 16 months of age by 1.8-fold and 2.0-fold when compared to NTg and TDP-43WT mice respectively (blue bar compared to orange and green bars in Fig. 5g; n = 5, ***P < 0.001). C5aR1 mRNA expression was also significantly increased by 1.7-fold and 1.6-fold at 10 months of age and by 1.7-fold and 2.0-fold at 16 months of age when compared to NTg and TDP-43WT controls, respectively (blue bar compared to orange and green bars in Fig. 5h; n = 5, *P < 0.05, **P < 0.01, ***P < 0.001). Taken together, these results suggest that dysregulation of the complement system also occurs in the TA muscle of TDP-43Q331K mice, which could contribute to the disease pathology in these animals.
Fig. 5
Fig. 5

Dysregulation of complement components in tibialis anterior muscle of TDP-43Q331K mice at three different ages of disease progression. a–f The mRNA expression profiles of the following complement components: C1qB (a, classical pathway), C4 (b, classical/lectin pathway), fB (c, alternative pathway), C3 (d, central component), CD55 (e, regulator) and CD59a (f, regulator) in tibialis anterior muscle of TDP-43Q331K mice (blue bars) relative to non-transgenic (NTg, orange bars) and TDP-43WT (green bars) mice during 3, 10 and 16 months of age. g The protein expression of C5a in the tibialis anterior muscle of NTg (orange bars), TDP-43WT (green bars) and TDP-43Q331K (blue bars) mice at 16 months. h mRNA expression of C5aR1 in the tibialis anterior muscle of TDP-43Q331K mice relative to age-matched NTg and TDP-43WT mice at three different ages. Data are expressed as means ± SEM (n = 5 mice/group, *P < 0.05, **P < 0.01, ***P < 0.001, one-way ANOVA with Tukey’s post hoc test for each age)

Motor dysfunction in TDP-43Q331K mice strongly correlates with lumbar spinal cord levels of C1qb, C3 and C5aR1

We next examined if there was any correlation between major complement transcript levels to the hind-limb grip strength of TDP-43Q331K mice. To investigate this aspect, we performed a Pearson correlation to measure the strength of the linear relationship between C1qB, C3 and C5aR1 mRNA levels in the lumbar spinal cord and TA muscle of NTg, TDP-43WT and TDP-43Q331K mice with their hind-limb grip strength. We found a strong negative correlation between lumbar spinal cord mRNA expression of C1qB (r = − 0.6824), C3 (r = − 0.8282) and C5aR1 (r = − 0.7428) to hind-limb grip strength of these animals, with C3 presenting the strongest correlation (n = 30, ****P < 0.0001; Fig. 6a–c). Furthermore, a moderate to strong negative correlation between TA mRNA levels of C1qB (r = − 0.6624) and C5aR1 (r = − 0.4116) to hind-limb grip strength was observed, whereas a moderate positive correlation was identified between C3 mRNA levels (r = 0.4149) to hind-limb grip strength (n = 25, *P < 0.05, ***P < 0.001; Fig. 6d–f). These results indicate that changes in complement transcript levels in the spinal cord and TA muscle directly correlates with the decrease in hind-limb grip strength (i.e. increased in ALS symptoms) in this ALS model.
Fig. 6
Fig. 6

Motor dysfunction in TDP-43Q331K mice strongly correlates with lumbar spinal cord/tibialis anterior muscle levels of C1qB, C3 and C5aR1. A strong negative correlation between lumbar spinal cord mRNA expression of C1qB (a, r = − 0.6824), C3 (b, r = − 0.8282) and C5aR1 (c, r = − 0.7428) with hind-limb grip strength of non-transgenic (NTg), TDP-43WT and TDP-43Q331K mice, where C3 demonstrated the strongest correlation (n = 30, ****P < 0.0001, Pearson correlation). A moderate to strong negative correlation between tibialis anterior muscle mRNA levels of C1qB (d, r = − 0.6624) and C5aR1 (f, r = − 0.4116) with hind-limb grip strength of NTg, TDP-43WT and TDP-43Q331K mice, whereas a moderate positive correlation between C3 mRNA levels (e, r = 0.4149) was observed (n = 25, *P < 0.05, ***P < 0.001, Pearson correlation). Pearson correlation coefficient was used to measure the strength of the linear relationship between complement mRNA expression and hind-limb grip strength

Discussion

The major findings of the current study are that components of the complement system are dysregulated in a transgenic mutant TDP43-based mouse model of ALS. It has now been well documented that the complement cascade is synthesised by neurons, astrocytes and microglia within the central nervous system and is involved in the disease progression of ALS, with evidence from both human patients and rodent models [8]. The present study further adds to this knowledge, demonstrating alteration of mRNA expression in major complement factors including C1qB, C4, fB, C3, C5a, C5aR1 and regulator CD55 in the spinal cord and TA muscles of TDP-43Q331K mice, which is suggestive of a progressive dysregulation of complement in this model. These results are similar to our previous demonstration of complement dysregulation in the spinal cord and TA muscle of hSOD1G93A mice [11, 15], indicating that complement activation occurs in response to motor neuron death and muscle denervation regardless of which ALS-related gene mutation is present. Furthermore, complement activation, and subsequent C5aR1 activation, could be a common mechanism of pathology in most forms of ALS.

The present study provided evidence for the dysregulation of classical/lectin, alternative and central component to all pathways of the complement system in the spinal cord and TA muscle of TDP-43Q331K mice during ALS disease progression. This is in line with numerous studies including our own, where increased C1qB, C4 and C3 was found in the spinal cord of hSOD1G93A mice [11, 15, 22, 23]. This upregulation of C1qB and C3 in the spinal cord could assist in the removal of dying motor neurons via opsonisation through microglia activation during disease progression in TDP-43Q331K mice, similar to what is evident in hSOD1G93A mice [15]. Similarly, we identified upregulation of C1qB, C4 and fB in the TA muscles of TDP-43Q331K mice, however, found downregulation of C3 in the TA muscles of TDP-43Q331K mice, which is contradictory of what is shown in hSOD1G93A mice. The upregulation of C1qB in the TA muscles could assist in the removal of degenerating neuromuscular synapses via phagocytosis during disease progression in TDP-43Q331K mice, similar to what has been shown in hSOD1G93A mice [22]. The reason for the differential expression of C3 in the TA muscle of TDP-43Q331K mice compared with hSOD1G93A mice is unclear, but could be due to the difference in the severity of the disease with TDP-43Q331K mice showing milder ALS-like symptoms compared to hSOD1G93A mice [13, 14].

We additionally showed decreased mRNA expression levels of CD55 in the spinal cord and increased expression levels in TA muscle, which suggests that homeostatic balance of the complement system may be perturbed in TDP-43Q331K mice, leading to the over activation of complement system. These findings support other studies, which have also shown decreased CD55 mRNA levels in the spinal cord, with deficiency in CD55 exacerbating neurodegeneration [15, 22, 24]. Our findings also support previous studies identifying increased CD55 expression in intercostal muscles of ALS patients and TA muscles of hSOD1G93A mice [11, 25]. Upregulation of CD55 at the motor end-plates of ALS patients and hSOD1G93A mice could be a mechanism to protect against high levels of complement activation at the neuromuscular junction. Interestingly, the current study did not demonstrate any changes in the other regulator CD59a in the spinal cord or TA muscle of TDP-43Q331K mice. This is in contrast to previous studies, where CD59a was decreased in the spinal cord of hSOD1G93A mice, while it increased in the intercostal muscle of ALS patients and TA muscle of hSOD1G93A mice [11, 15, 25]. The difference in the expression changes between different models and patients could be attributed to the severity of the disease, as TDP-43Q331K mice show milder ALS-like symptoms compared to hSOD1G93A mice [9, 13–15]. However, further investigation is required into the expression and localisation of membrane attack complex in TDP-43Q331K mice, and its putative correlation with motor neuron loss. Regardless, collectively our data adds further support to the notion that dysregulated complement activity may play an important role in accelerating motor neuron loss and neuromuscular junction denervation, ultimately driving the progression of ALS.

Among the complement activation effector molecules, C5a is considered the most potent peptide, with its signalling through its main receptor C5aR1 having detrimental effects in multiple neurodegenerative diseases, including ALS [9, 10, 12, 26, 27]. To obtain a better understanding of its role in disease progression of ALS, we analysed C5a and its receptor C5aR1 in the spinal cord and TA muscle of TDP-43Q331K mice. C5a protein levels in the spinal cord did not change between NTg, TDP-43WT and TDP-43Q331K mice, whereas it increased in the TA muscles of TDP-43Q331K mice. This is in line with C5a protein levels at mid-symptomatic stage in the spinal cord and TA muscle of hSOD1G93A mice [11], suggesting that enhanced C5a-C5aR1 signalling may affect disease progression of ALS, similar to what has been found in hSOD1G93A mice. This finding is interesting given that C3 mRNA transcript levels were higher in the spinal cord compared to TA muscle in these animals. The reason for the increase in muscle C5a, despite reduced C3 transcript expression is unclear; however, it should be noted that mRNA expression does not directly equate with the degree of complement activation. Furthermore, there is a well-described C3-bypass (extrinsic) complement activation pathway that enables C3-independent cleavage of C5 [28]. Thus, C5a protein levels do not necessarily correlate with C3 mRNA expression within a tissue.

Numerous studies have demonstrated upregulation of C5aR1 within the spinal cord and TA muscles of hSOD1G93A rats and mice, as well as human ALS patients, suggesting that heightened C5a-C5aR1 signalling plays a role in ALS pathology [9–12, 15, 29]. In the present study, we demonstrated that the mRNA expression of C5aR1 in TDP-43Q331K spinal cord and TA muscles is elevated, confirming dysregulation of downstream terminal complement pathway in these animals. This suggests that enhanced C5a-C5aR1 signalling may affect the disease progression of ALS in both spinal cord and TA muscles of TDP-43Q331K mice, similar to what is evident in hSOD1G93A mice [9, 11, 15]. In addition to an increase in C5aR1 in TDP-43Q331K mice, the present study also revealed C5aR1 on motor neurons and microglia in NTg, TDP-43WT and TDP-43Q331K mice. This is in concordance with previous studies showing C5aR1 on motor neurons and microglia in other mouse models of ALS [15, 20], indicating that C5a-C5aR1 signalling could play a role in facilitating microglia activation and motor neuron death regardless of which ALS-related gene mutation is present.

In line with increased complement components in TDP-43Q331K mice, the present study also revealed a strong correlation between C1qB, C3 and C5aR1 mRNA levels in the spinal cord and TA muscles to the hind-limb grip strength of these animals, suggesting that the mRNA levels of complement system could be a direct correlate of motor neuron loss and neuromuscular junction denervation throughout disease progression, that we and others have previously demonstrated in this mouse model [13, 14]. These finding are consistent with increased astrocyte and microglia numbers/activation in these animals, indicating that neuroinflammation could be a good indicator of disease severity and pathology.

Conclusions

In summary, the current study has demonstrated upregulation of major complement factors, together with decreased levels of the negative complement regulator CD55, in TDP-43Q331K mice. This suggests that complement activation and/or its dysregulation could play an important role in motor neuron loss and neuromuscular junction denervation in this TDP43-based mouse model of ALS. Expression of the C5a receptor, C5aR1, was also upregulated in TDP-43Q331K mice, predominantly due to increased microglial/macrophage C5aR1 expression, and was strongly correlated with resulting motor decline. Taken together, these results indicate that heightened complement activation and enhanced C5aR1 signalling may play a crucial role in pathophysiology of the TDP-43Q331K ALS model, further validating C5aR1 as a potential therapeutic target for all forms of ALS.

Abbreviations

ALS: 

Amyotrophic lateral sclerosis

BSA: 

Bovine serum albumin

DAPI: 

Diamidino-2-phenylindole

DS: 

Donkey serum

NTg: 

Non-transgenic

TA: 

Tibialis anterior

Declarations

Acknowledgements

The authors would like to sincerely thank Associate Professor Massimo A. Hilliard for carefully going through the manuscript and giving us feedback to improve our manuscript. We also thank Maryam Shayegh for her technical support with genotyping mice, Mary White for general laboratory support and Dr. Shaun Walters for his support with microscopy.

Funding

JDL holds a Motor Neuron Disease Research Institute of Australia (MNDRIA) Postdoctoral Fellowship. TMW is supported by a NHMRC Career Development Fellowship. The research was funded by grants from the MNDRIA to PGN (GIA1429 and GIA1547), TMW (GIA1865) and JDL (GIA1830) and a National Health and Medical Research Council (NHMRC) Project grant (APP1082271) to TMW.

Authors’ contributions

JDL, TMW and PGN conceived the project. JDL, TMW and PGN designed the study. JDL, SCL, EFW and RL performed the experiments in the laboratories of PGN and TMW. All authors contributed to the analyses and/or interpreted the data. JDL wrote the paper with contributions from TMW and PGN. All authors read and approved the final manuscript.

Ethics approval

All experimental procedures were approved by the University of Queensland Animal Ethics Committee and complied with the policies and regulations regarding animal experimentation. They were conducted in accordance with the Queensland Government Animal Research Act 2001, associated Animal Care and Protection Regulations (2002 and 2008) and the Australian Code of Practice for the Care and Use of Animals for Scientific Purposes, 8th Edition (National Health and Medical Research Council, 2013). ARRIVE guidelines have been followed in the preparation of the manuscript.

Competing interests

The authors declare that they have no competing interests.

Publisher’s Note

Springer Nature remains neutral with regard to jurisdictional claims in published maps and institutional affiliations.

Open AccessThis article is distributed under the terms of the Creative Commons Attribution 4.0 International License (http://creativecommons.org/licenses/by/4.0/), which permits unrestricted use, distribution, and reproduction in any medium, provided you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons license, and indicate if changes were made. The Creative Commons Public Domain Dedication waiver (http://creativecommons.org/publicdomain/zero/1.0/) applies to the data made available in this article, unless otherwise stated.

Authors’ Affiliations

(1)
School of Biomedical Sciences, the University of Queensland, St Lucia, Brisbane, QLD, 4072, Australia
(2)
University of Queensland Centre for Clinical Research, the University of Queensland, Herston, Brisbane, QLD, 4029, Australia
(3)
Queensland Brain Institute, the University of Queensland, St Lucia, Brisbane, QLD, 4072, Australia

References

  1. DeJesus-Hernandez M, Mackenzie IR, Boeve BF, Boxer AL, Baker M, Rutherford NJ, Nicholson AM, Finch NA, Flynn H, Adamson J, et al. Expanded GGGGCC hexanucleotide repeat in noncoding region of C9ORF72 causes chromosome 9p-linked FTD and ALS. Neuron. 2011;72:245–56.View ArticlePubMedPubMed CentralGoogle Scholar
  2. Kwiatkowski TJ Jr, Bosco DA, Leclerc AL, Tamrazian E, Vanderburg CR, Russ C, Davis A, Gilchrist J, Kasarskis EJ, Munsat T, et al. Mutations in the FUS/TLS gene on chromosome 16 cause familial amyotrophic lateral sclerosis. Science. 2009;323:1205–8.View ArticlePubMedGoogle Scholar
  3. Rosen DR, Siddique T, Patterson D, Figlewicz DA, Sapp P, Hentati A, Donaldson D, Goto J, O'Regan JP, Deng H-X. Mutations in Cu/Zn superoxide dismutase gene are associated with familial amyotrophic lateral sclerosis. Nature. 1993;362:59–62.View ArticlePubMedGoogle Scholar
  4. Sreedharan J, Blair IP, Tripathi VB, Hu X, Vance C, Rogelj B, Ackerley S, Durnall JC, Williams KL, Buratti E, et al. TDP-43 mutations in familial and sporadic amyotrophic lateral sclerosis. Science. 2008;319:1668–72.View ArticlePubMedGoogle Scholar
  5. Williams KL, Warraich ST, Yang S, Solski JA, Fernando R, Rouleau GA, Nicholson GA, Blair IP. UBQLN2/ubiquilin 2 mutation and pathology in familial amyotrophic lateral sclerosis. Neurobiol Aging. 2012;33:2527 e2523–10.Google Scholar
  6. Taylor JP, Brown RH Jr, Cleveland DW. Decoding ALS: from genes to mechanism. Nature. 2016;539:197–206.View ArticlePubMedPubMed CentralGoogle Scholar
  7. Woodruff TM, Costantini KJ, Taylor SM, Noakes PG. Role of complement in motor neuron disease: animal models and therapeutic potential of complement inhibitors. Adv Exp Med Biol. 2008;632:143–58.PubMedGoogle Scholar
  8. Brennan FH, Lee JD, Ruitenberg MJ, Woodruff TM. Therapeutic targeting of complement to modify disease course and improve outcomes in neurological conditions. Semin Immunol. 2016;28:292–308.View ArticlePubMedGoogle Scholar
  9. Lee JD, Kumar V, Fung JN, Ruitenberg MJ, Noakes PG, Woodruff TM. Pharmacological inhibition of complement C5a-C5a1 receptor signalling ameliorates disease pathology in the hSOD1(G93A) mouse model of amyotrophic lateral sclerosis. Br J Pharmacol. 2017;174:689–99.View ArticlePubMedPubMed CentralGoogle Scholar
  10. Woodruff TM, Lee JD, Noakes PG. Role for terminal complement activation in amyotrophic lateral sclerosis disease progression. Proc Natl Acad Sci U S A. 2014;111:E3–4.View ArticlePubMedGoogle Scholar
  11. Wang HA, Lee JD, Lee KM, Woodruff TM, Noakes PG. Complement C5a-C5aR1 signalling drives skeletal muscle macrophage recruitment in the hSOD1(G93A) mouse model of amyotrophic lateral sclerosis. Skelet Muscle. 2017;7:10.View ArticlePubMedPubMed CentralGoogle Scholar
  12. Woodruff TM, Costantini KJ, Crane JW, Atkin JD, Monk PN, Taylor SM, Noakes PG. The complement factor C5a contributes to pathology in a rat model of amyotrophic lateral sclerosis. J Immunol. 2008;181:8727–34.View ArticlePubMedGoogle Scholar
  13. Arnold ES, Ling SC, Huelga SC, Lagier-Tourenne C, Polymenidou M, Ditsworth D, Kordasiewicz HB, McAlonis-Downes M, Platoshyn O, Parone PA, et al. ALS-linked TDP-43 mutations produce aberrant RNA splicing and adult-onset motor neuron disease without aggregation or loss of nuclear TDP-43. Proc Natl Acad Sci U S A. 2013;110:E736–45.View ArticlePubMedPubMed CentralGoogle Scholar
  14. Chand KK, Lee KM, Lee JD, Qiu H, Willis EF, Lavidis NA, Hilliard MA, Noakes PG. Defects in synaptic transmission at the neuromuscular junction precedes motor deficits in a TDP-43(Q331K) transgenic mouse model of amyotrophic lateral sclerosis. FASEB J. 2018;32:2676–89.Google Scholar
  15. Lee JD, Kamaruzaman NA, Fung JN, Taylor SM, Turner BJ, Atkin JD, Woodruff TM, Noakes PG. Dysregulation of the complement cascade in the hSOD1G93A transgenic mouse model of amyotrophic lateral sclerosis. J Neuroinflammation. 2013;10:119.View ArticlePubMedPubMed CentralGoogle Scholar
  16. Lee JY, Lee JD, Phipps S, Noakes PG, Woodruff TM. Absence of toll-like receptor 4 (TLR4) extends survival in the hSOD1 G93A mouse model of amyotrophic lateral sclerosis. J Neuroinflammation. 2015;12:90.View ArticlePubMedPubMed CentralGoogle Scholar
  17. Watson C, Paxinos G, Kayalioglu G. The spinal cord. 1st ed. London: Academic Press; 2009.Google Scholar
  18. Li Q, Barres BA. Microglia and macrophages in brain homeostasis and disease. Nat Rev Immunol. 2017;18:225–42.Google Scholar
  19. Philips T, Robberecht W. Neuroinflammation in amyotrophic lateral sclerosis: role of glial activation in motor neuron disease. Lancet Neurol. 2011;10:253–63.View ArticlePubMedGoogle Scholar
  20. Humayun S, Gohar M, Volkening K, Moisse K, Leystra-Lantz C, Mepham J, McLean J, Strong MJ. The complement factor C5a receptor is upregulated in NFL−/− mouse motor neurons. J Neuroimmunol. 2009;210:52–62.View ArticlePubMedGoogle Scholar
  21. Woodruff TM, Ager RR, Tenner AJ, Noakes PG, Taylor SM. The role of the complement system and the activation fragment C5a in the central nervous system. NeuroMolecular Med. 2010;12:179–92.View ArticlePubMedGoogle Scholar
  22. Heurich B, El Idrissi NB, Donev RM, Petri S, Claus P, Neal J, Morgan BP, Ramaglia V. Complement upregulation and activation on motor neurons and neuromuscular junction in the SOD1 G93A mouse model of familial amyotrophic lateral sclerosis. J Neuroimmunol. 2011;235:104–9.View ArticlePubMedGoogle Scholar
  23. Lobsiger CS, Boillee S, Pozniak C, Khan AM, McAlonis-Downes M, Lewcock JW, Cleveland DW. C1q induction and global complement pathway activation do not contribute to ALS toxicity in mutant SOD1 mice. Proc Natl Acad Sci U S A. 2013;110:E4385–92.View ArticlePubMedPubMed CentralGoogle Scholar
  24. Wang Y, Li Y, Dalle Lucca SL, Simovic M, Tsokos GC, Dalle Lucca JJ. Decay accelerating factor (CD55) protects neuronal cells from chemical hypoxia-induced injury. J Neuroinflammation. 2010;7:24.View ArticlePubMedPubMed CentralGoogle Scholar
  25. Bahia El Idrissi N, Bosch S, Ramaglia V, Aronica E, Baas F, Troost D. Complement activation at the motor end-plates in amyotrophic lateral sclerosis. J Neuroinflammation. 2016;13:72.View ArticlePubMedPubMed CentralGoogle Scholar
  26. Fonseca MI, Ager RR, Chu SH, Yazan O, Sanderson SD, LaFerla FM, Taylor SM, Woodruff TM, Tenner AJ. Treatment with a C5aR antagonist decreases pathology and enhances behavioral performance in murine models of Alzheimer's disease. J Immunol. 2009;183:1375–83.View ArticlePubMedPubMed CentralGoogle Scholar
  27. Woodruff TM, Crane JW, Proctor LM, Buller KM, Shek AB, de Vos K, Pollitt S, Williams HM, Shiels IA, Monk PN, Taylor SM. Therapeutic activity of C5a receptor antagonists in a rat model of neurodegeneration. FASEB J. 2006;20:1407–17.View ArticlePubMedGoogle Scholar
  28. Huber-Lang M, Younkin EM, Sarma JV, Riedemann N, McGuire SR, Lu KT, Kunkel R, Younger JG, Zetoune FS, Ward PA. Generation of C5a by phagocytic cells. Am J Pathol. 2002;161:1849–59.View ArticlePubMedPubMed CentralGoogle Scholar
  29. Mantovani S, Gordon R, Macmaw JK, Pfluger CM, Henderson RD, Noakes PG, McCombe PA, Woodruff TM. Elevation of the terminal complement activation products C5a and C5b-9 in ALS patient blood. J Neuroimmunol. 2014;276:213–8.View ArticlePubMedGoogle Scholar

Copyright

© The Author(s). 2018

Advertisement

Journal of Neuroinflammation

ISSN: 1742-2094