The Microglial Trem2 R47H Alzheimer’s Disease Risk Variant Impairs Early Hippocampal Synaptic Remodeling

The Microglial Trem2 R47H Alzheimer’s Disease Risk Variant Impairs Early Hippocampal Synaptic Remodeling | bioRxiv /* */ /* */ <!-- <!-- /*! * yepnope1.5.4 * (c) WTFPL, GPLv2 */ (function(a,b,c){function d(a){return"[object Function]"==o.call(a)}function e(a){return"string"==typeof a}function f(){}function g(a){return!a||"loaded"==a||"complete"==a||"uninitialized"==a}function h(){var a=p.shift();q=1,a?a.t?m(function(){("c"==a.t?B.injectCss:B.injectJs)(a.s,0,a.a,a.x,a.e,1)},0):(a(),h()):q=0}function i(a,c,d,e,f,i,j){function k(b){if(!o&&g(l.readyState)&&(u.r=o=1,!q&&h(),l.onload=l.onreadystatechange=null,b)){"img"!=a&&m(function(){t.removeChild(l)},50);for(var d in y[c])y[c].hasOwnProperty(d)&&y[c][d].onload()}}var j=j||B.errorTimeout,l=b.createElement(a),o=0,r=0,u={t:d,s:c,e:f,a:i,x:j};1===y[c]&&(r=1,y[c]=[]),"object"==a?l.data=c:(l.src=c,l.type=a),l.width=l.height="0",l.onerror=l.onload=l.onreadystatechange=function(){k.call(this,r)},p.splice(e,0,u),"img"!=a&&(r||2===y[c]?(t.insertBefore(l,s?null:n),m(k,j)):y[c].push(l))}function j(a,b,c,d,f){return q=0,b=b||"j",e(a)?i("c"==b?v:u,a,b,this.i++,c,d,f):(p.splice(this.i++,0,a),1==p.length&&h()),this}function k(){var a=B;return a.loader={load:j,i:0},a}var l=b.documentElement,m=a.setTimeout,n=b.getElementsByTagName("script")[0],o={}.toString,p=[],q=0,r="MozAppearance"in l.style,s=r&&!!b.createRange().compareNode,t=s?l:n.parentNode,l=a.opera&&"[object Opera]"==o.call(a.opera),l=!!b.attachEvent&&!l,u=r?"object":l?"script":"img",v=l?"script":u,w=Array.isArray||function(a){return"[object Array]"==o.call(a)},x=[],y={},z={timeout:function(a,b){return b.length&&(a.timeout=b[0]),a}},A,B;B=function(a){function b(a){var a=a.split("!"),b=x.length,c=a.pop(),d=a.length,c={url:c,origUrl:c,prefixes:a},e,f,g;for(f=0;f<d;f++)g=a[f].split("="),(e=z[g.shift()])&&(c=e(c,g));for(f=0;f<b;f++)c=x[f](c);return c}function g(a,e,f,g,h){var i=b(a),j=i.autoCallback;i.url.split(".").pop().split("?").shift(),i.bypass||(e&&(e=d(e)?e:e[a]||e[g]||e[a.split("/").pop().split("?")[0]]),i.instead?i.instead(a,e,f,g,h):(y[i.url]?i.noexec=!0:y[i.url]=1,f.load(i.url,i.forceCSS||!i.forceJS&&"css"==i.url.split(".").pop().split("?").shift()?"c":c,i.noexec,i.attrs,i.timeout),(d(e)||d(j))&&f.load(function(){k(),e&&e(i.origUrl,h,g),j&&j(i.origUrl,h,g),y[i.url]=2})))}function h(a,b){function c(a,c){if(a){if(e(a))c||(j=function(){var a=[].slice.call(arguments);k.apply(this,a),l()}),g(a,j,b,0,h);else if(Object(a)===a)for(n in m=function(){var b=0,c;for(c in a)a.hasOwnProperty(c)&&b++;return b}(),a)a.hasOwnProperty(n)&&(!c&&!--m&&(d(j)?j=function(){var a=[].slice.call(arguments);k.apply(this,a),l()}:j[n]=function(a){return function(){var b=[].slice.call(arguments);a&&a.apply(this,b),l()}}(k[n])),g(a[n],j,b,n,h))}else!c&&l()}var h=!!a.test,i=a.load||a.both,j=a.callback||f,k=j,l=a.complete||f,m,n;c(h?a.yep:a.nope,!!i),i&&c(i)}var i,j,l=this.yepnope.loader;if(e(a))g(a,0,l,0);else if(w(a))for(i=0;i (function(w,d,s,l,i){w[l]=w[l]||[];w[l].push({'gtm.start':new Date().getTime(),event:'gtm.js'});var f=d.getElementsByTagName(s)[0];var j=d.createElement(s);var dl=l!='dataLayer'?'&l='+l:'';j.src='//www.googletagmanager.com/gtm.js?id='+i+dl;j.type='text/javascript';j.async=true;f.parentNode.insertBefore(j,f);})(window,document,'script','dataLayer','GTM-M677548'); Skip to main content Home About Submit ALERTS / RSS Search for this keyword Advanced Search New Results The Microglial Trem2 R47H Alzheimer’s Disease Risk Variant Impairs Early Hippocampal Synaptic Remodeling View ORCID Profile Abdulaziz Aljawder , View ORCID Profile Damian M. Cummings , View ORCID Profile Jack Wood , Frances A. Edwards doi: https://doi.org/10.1101/2025.08.22.671714 Abdulaziz Aljawder 1 Department of Neuroscience, Physiology & Pharmacology, University College London , Gower Street, London WC1E 6BT, United Kingdom 2 Department of Physiology, College of Medicine and Health Sciences, Arabian Gulf University , Block 329, Manama, Bahrain Find this author on Google Scholar Find this author on PubMed Search for this author on this site ORCID record for Abdulaziz Aljawder Damian M. Cummings 1 Department of Neuroscience, Physiology & Pharmacology, University College London , Gower Street, London WC1E 6BT, United Kingdom Find this author on Google Scholar Find this author on PubMed Search for this author on this site ORCID record for Damian M. Cummings Jack Wood 1 Department of Neuroscience, Physiology & Pharmacology, University College London , Gower Street, London WC1E 6BT, United Kingdom Find this author on Google Scholar Find this author on PubMed Search for this author on this site ORCID record for Jack Wood Frances A. Edwards 1 Department of Neuroscience, Physiology & Pharmacology, University College London , Gower Street, London WC1E 6BT, United Kingdom Find this author on Google Scholar Find this author on PubMed Search for this author on this site For correspondence: f.a.edwards{at}ucl.ac.uk Abstract Full Text Info/History Metrics Preview PDF Summary The rare R47H variant of the microglial TREM2 gene increases Alzheimer’s disease (AD) risk, but its effects on early hippocampal circuitry remain unclear. We examined basal synaptic transmission and synaptic density in 3-week-old Trem2 R47H knockin mice using in vitro whole-cell patch clamp recordings and immunohistochemistry. R47H mice displayed increased synaptic density in CA1 and CA3 regions, consistent with a trend towards increased spontaneous excitatory current frequency. Evoked synaptic currents, miniature EPSCs and spontaneous inhibitory current frequency were unaltered. Microglia shape synaptic circuits in early development by removing inactive synapses. Thus, inhibition of this role due to the Trem2 R47H mutation decreases the microglial reshaping of hippocampal connectivity during early postnatal development, without overtly altering basal synaptic activity. With ongoing synaptic plasticity, such early structural changes may predispose neural circuits to later dysfunction, particularly in the context of AD pathology, highlighting the importance of microglial TREM2 in developmental synaptic refinement. Introduction Triggering receptor expressed on myeloid cells 2 (TREM2) is a microglial cell surface receptor predominantly expressed in the central nervous system 1 , 2 . Homozygosity for the loss-of-function variants in the TREM2 gene causes Nasu-Hakola disease, which is characterized by early-onset dementia, multifocal bone cysts, and may also lead to epilepsy 3 - 5 . Additionally, a range of experimental approaches have identified that heterozygosity for the rare R47H missense variant in TREM2 as one of the strongest genetic risk factors for sporadic Alzheimer’s disease (AD), conferring up to a fourfold increase in disease risk 6 - 11 . During development, TREM2, together with neuronal activity and complement proteins such as C1q, C3, and CR3, plays an essential role in pruning inactive synapses, a process critical for the maturation and refinement of functional neuronal circuits 12 - 14 . Considering the essential role of TREM2 in development and maintenance of synaptic networks, loss of function mutations in TREM2 might be expected to result in developmental differences in such networks, possibly contributing to the risk of AD in later life. In a recent study in 5-6-month-old mice heterozygous for the Trem2 R47H mutation, Das et al. (2023) reported changes in synaptic networks in both wild type (WT) mice and in a rapidly progressing model of early AD ( App NL-G-F/NL-G-F knockin mice) 15 , 16 . Specifically, the Trem2 R47H expression increased susceptibility to kainate-induced seizure activity, which the authors attributed to elevated synapse density observed in the cortex of both WT and App knockin mice carrying the Trem2 mutation. No significant difference was observed in synaptic density in the hippocampus at this age 15 . In AD, Trem2 expression is upregulated in microglia in areas where plaques are abundant 17 , particularly in microglia directly in contact with plaques 18 . Increased expression of Trem2 leads to an anti-inflammatory and pro-phagocytic phenotype in microglia 18 , resulting in an increased clearance of damaged synapses 19 , 20 . However, in the presence of the Trem2 R47H mutation, microglial binding with its ligands might be dysregulated, which could result in impaired phagocytosis 21 . Gratuze et al. reported that the Trem2 R47H variant attenuates the loss of synapses in 9-month-old PS19 tauopathy mice expressing human tau with the P301S mutation 22 . Despite growing evidence for synaptic and network modulation impairment, research on the electrophysiological impact of the Trem2 R47H variant remains limited. Studies on a different Trem2 loss-of-function mutation, Y38C , in 6-month-old mice, have demonstrated a significant impairment in long-term potentiation (LTP) 23 . In contrast, in Trem2 knockout mice, basal neurotransmission and LTP were found unaffected at 6 months of age, whereas 19-month-old mice exhibited enhanced LTP, despite unchanged basal transmission and paired-pulse ratios 24 . In pre-weaned (P18-20) Trem2 knockout mice, increased synaptic density and elevated frequencies of miniature excitatory postsynaptic currents (mEPSCs) have been reported, suggesting that loss of TREM2 function during early development can promote neuronal hyperexcitability 14 . Moreover, the same study reported that Trem2 knockout resulted in reduced functional connectivity between the hippocampus and prefrontal cortex, accompanied by abnormalities in social behavior 14 . Early-life events can influence later susceptibility to neurodegenerative disorders (reviewed in 25 , 26 ). Studies of such susceptibility to early-life influence are often related to changes in immune function with most reports relating to environmental factors such as perinatal hypoxia 27 , disturbance of the microbiome 28 or other environmental factors 29 . In contrast, genetic variants that increase the risk of AD have usually been studied in terms of their role in directly exacerbating the progression of disease in later life. However, dysfunctions caused by genetic variants, for example in microglia, are present throughout life and so may well increase the vulnerability of an individual to AD due to early-life effects in addition to later exacerbation of disease progression. As microglia have an important role in refining synaptic circuits in early development, understanding how genetic variants, such as TREM2 R47H, affect microglia-mediated synaptic development may provide crucial insights into additional mechanisms by which this variant increases AD risk. In this study, using immunohistochemistry, we demonstrate an increased density of synapses in both the CA1 and CA3 subfields of the hippocampus. Whole-cell patch clamp recordings show a trend towards increased frequency of spontaneous excitatory postsynaptic currents (sEPSCs), possibly reflecting a change in excitatory/inhibitory balance, but overall, there is little change to basal excitatory or inhibitory synaptic activity in CA1 pyramidal neurons of 18–22-day-old mice. This may reflect the fact that synapses normally pruned in development are those with little activity and so their pathological retention is not detectable in electrophysiological recordings. However, as ongoing synaptic plasticity occurs over time, additional synapses may be a source of hyperactivity that could contribute to vulnerability to AD in older age. Results In order to assess the effects of the Trem2 R47H mutation on synaptic function in young mice, whole-cell patch clamp recordings were performed from CA1 pyramidal neurons in acute hippocampal slices prepared from mice homozygous for the Trem2 R47H variant and WT controls at 18–22 days old. Synaptic activity was analyzed by quantifying the frequency, amplitude, and time course of sEPSCs and mEPSCs. In addition, evoked synaptic responses were recorded using paired-pulse stimulation of the Schaffer collateral pathway to calculate paired-pulse ratios, a measure inversely related to neurotransmitter release probability 30 . Trem2 R47H variant does not alter spontaneous inhibitory synaptic currents Spontaneous synaptic activity recorded in artificial cerebrospinal fluid (ACSF) reflects a mixture of sEPSCs and spontaneous inhibitory postsynaptic currents (sIPSCs). To quantify the relative contribution of sEPSCs, we compared the mean frequency of total spontaneous events in WT recordings (4.23 Hz) with the mean frequency of pharmacologically isolated sEPSCs (0.074 Hz). sEPSCs represented less than 2% of the total events, indicating that approximately 98% of the spontaneous currents recorded under baseline conditions were inhibitory. Therefore, we refer to the composite signal as sIPSCs ( Figure 1 ). Download figure Open in new tab Figure 1. The Trem2 R47H variant does not affect spontaneous inhibitory postsynaptic currents. (A) Representative sIPSC current traces recorded from CA1 pyramidal neurons in WT (black) and Trem2 R47H (blue) mice. Scale bars are indicated. (B) Extended 10 s traces of sIPSCs in WT and Trem2 R47H neurons. (C–E) Quantification of sIPSC frequency (C), median amplitude (D), and τ (E) across genotypes. Data are presented as mean + SEM. Statistical comparisons were performed using unpaired t -tests. Analysis of sIPSCs in CA1 pyramidal neurons from Trem2 R47H and WT mice (P18–22) revealed no significant differences between genotypes in event frequency, median amplitude, or decay time-constant τ, suggesting that the Trem2 R47H variant does not impact basal inhibitory synaptic transmission ( Figure 1 ). Trem2 R47H variant tends to increase sEPSC but not mEPSC frequency To isolate sEPSCs, GABA A receptor-mediated inhibition was pharmacologically blocked using gabazine (6 μM, Figure 2A ). Analysis of sEPSCs recorded from CA1 pyramidal neurons revealed that event frequency in Trem2 R47H mice tended to be slightly higher than in WT controls (P = 0.07; Figure 2B–C ). There were no significant differences between genotypes in either the median amplitude or τ of sEPSCs ( Figure 2D–E ). Download figure Open in new tab Figure 2. The Trem2 R47H variant tends to increase the frequency of sEPSCs without affecting mEPSC. (A) Representative sEPSC traces from CA1 pyramidal neurons in WT (black) and Trem2 R47H (blue) mice. Scale bars are indicated. (B) 10 s continuous recordings of sEPSCs from WT and Trem2 R47H mice. (C–E) Quantification of sEPSC frequency (C), median amplitude (D), and τ (E). (F–H) Quantification of mEPSC frequency (F), median amplitude (G), and τ (H). Data are presented as mean + SEM. Statistical comparisons were performed using unpaired t -tests. To isolate mEPSCs, 1 μM tetrodotoxin was added to the extracellular solution containing gabazine, thereby blocking action potential-dependent synaptic activity. Analysis of mEPSCs recorded from CA1 pyramidal neurons revealed no significant differences between Trem2 R47H and WT mice in event frequency, amplitude, or τ (P > 0.05; Figure 2F-H ). Together, these findings suggest that the Trem2 R47H mutation tends to induce a small increase in sEPSC frequency, without affecting mEPSCs. R47H mutation does not alter the probability of glutamate release To assess potential changes in presynaptic glutamate release probability, we recorded evoked excitatory postsynaptic currents from CA1 pyramidal neurons in response to paired-pulse stimulation of the Schaffer collateral pathway at three different interstimulus intervals. The paired-pulse ratio is inversely related to neurotransmitter release probability ( Figure 3 ) 30 . Download figure Open in new tab Figure 3. The Trem2 R47H variant does not alter presynaptic glutamate release probability. (A) Representative traces of evoked excitatory postsynaptic currents recorded from CA1 pyramidal neurons in WT;(black) and Trem2 R47H (blue) mice at each inter-stimulus interval. Stimulus artifacts are attenuated for clarity. Scale bar is indicated. (B) Summary of paired-pulse ratios across all inter-stimulus intervals (25, 50, and 100 ms). (mean ± SEM). (C–E) Quantification of paired-pulse ratios at each inter-stimulus interval: 25 ms (C), 50 ms (D), and 100 ms (E). Data are presented as mean + SEM. Statistical comparisons in panel B were performed using two-way ANOVA, and panels C–E using unpaired t -tests. Quantification of paired-pulse ratios revealed no significant differences between Trem2 R47H and WT mice at any inter-stimulus interval tested ( Figure 3 ). Additionally, two-way ANOVA indicated no significant interaction between genotype and inter-stimulus interval, suggesting that the Trem2 R47H variant does not impact presynaptic release dynamics in the developing hippocampus. Trem2 R47H enhances synaptic density in the hippocampus If the Trem2 R47H variant impairs appropriate synaptic pruning during development, it is surprising that this has little, if any, effect on basal synaptic transmission in young mice. However, considering synaptic pruning targets relatively inactive synapses, i.e. those with a low release probability, it is possible that retention of such synapses would be difficult to detect with patch clamp techniques. We thus went on to investigate whether any difference in the density of excitatory synapses could be observed using immunohistochemistry. To investigate synaptic density alterations in the hippocampus, fluorescent immunohistochemistry was performed on brain sections from the fixed hemisphere of WT and Trem2 R47H mice. Presynaptic terminals were labelled with anti-Bassoon antibodies, postsynaptic densities with anti-Homer1 antibodies, and cell nuclei with DAPI ( Figure 4A ). High-resolution Airyscan imaging followed by 3D reconstruction in Imaris software was used for synaptic quantification ( Figure 4B ). Download figure Open in new tab Figure 4. The Trem2 R47H variant increases synaptic density in the hippocampus of young mice. (A) Representative confocal Airyscan-processed images of regions of interest in the hippocampus showing immunofluorescent labelling of presynaptic Bassoon (magenta), postsynaptic Homer1 (yellow), and nuclear DAPI (blue). Scale bars are indicated. (B) Three-dimensional reconstructions of z-stack images generated using Imaris software. Scale bars are indicated. (C) Colocalized synaptic puncta identified by Imaris analysis. Ci shows a raw higher magnification confocal image of colocalized Homer1 and Bassoon puncta; Cii displays the same region following Imaris colocalization reconstruction. Scale bars are indicated. (D) Quantification of synaptic density (colocalized puncta per µm 3 ) in CA1 and CA3 regions. Statistical analysis was performed using two-way ANOVA to assess the effects of genotype and hippocampal subregion. * P < 0.05 (main effect of genotype). Data are presented as mean ± SEM. Synaptic puncta were defined as Bassoon and Homer1 colocalized signals. Signals located within DAPI-labeled nuclear volumes were excluded from analysis to focus on extranuclear fluorescent signals ( Figure 4C ). Two-way ANOVA revealed a significant main effect of genotype, with Trem2 R47H mice exhibiting increased colocalized puncta density compared to WT controls ( Figure 4D ). No significant effects of region (CA1 versus CA3) or genotype-region interaction were observed ( Figure 4D ). These results indicate that the Trem2 R47H variant is associated with increased synaptic density, irrespective of hippocampal region. Discussion Using immunohistochemistry, this study indicates that the Trem2 R47H variant increases synaptic density in the CA1 and CA3 regions of the hippocampus during early postnatal development in mice. Although not reaching significance, the strong trend (P = 0.07) towards an increase in frequency of excitatory synaptic currents in the electrophysiological recordings is compatible with this observation. Mutations in TREM2 are well-established risk factors for sporadic AD 6 - 11 . At later stages, the R47H variant has been shown to attenuate plaque-associated upregulation of TREM2, and to exacerbate the accumulation of dystrophic neurites in AD pathology 19 , 31 . However, the mutation is present throughout development, and little is known about its effects on the initial establishment of the synaptic network. It seems likely that early effects will be present considering the critical role of TREM2 in the regulation of synaptic pruning 12 , 14 . It is thus not surprising that, in the presence of a mutation that decreases this tightly regulated pruning, an excess of synapses may persist, potentially disrupting synaptic homeostasis and leading to aberrant neuronal signaling. The fact that miniature excitatory current frequency was not changed could be counter to this observation. However, considering pruning in development preferentially removes synapses with low activity, this could initially mask the presence of these pathologically retained synapses in electrophysiological recordings in the absence of action potentials. In contrast, a previous study reported increased mEPSC frequency in Trem2 knockout mice at postnatal days 18–20, suggesting that knockout of the gene has a stronger effect than the mutation 14 . In our study, we found that the Trem2 R47H mutation does not alter the frequency, amplitude or decay time of sIPSCs, which reflect the net inhibitory synaptic input. Given our concurrent observation of increased excitatory synapse density and a trend toward increased sEPSC frequency in Trem2 R47H mutants, this suggests a change in the excitatory/inhibitory balance in CA1 pyramidal neurons. Moreover, with ongoing synaptic activity and potentiation over time, some, or all of these initially silent excitatory synapses could produce increasingly aberrant activity and indeed hyperactivity is a feature of early AD 32 . While we did not detect changes in inhibitory activity, consistent with the fact that the spine synapses studied in most reports of microglial pruning in early synapse development are largely excitatory, Favuzzi et al. demonstrated that, in the mouse somatosensory cortex, a specialized microglial subset expressing GABA B1 receptors selectively prunes inhibitory synapses during early postnatal development, supporting the possibility of synapse-type-specific microglial targeting 33 . It should be noted that the type of electrophysiological experiment reported in the present study records the net synaptic activity of many subtypes of interneurons received by pyramidal neurons, and such recordings are dominated by the basket cell inputs that are prominent on the soma of pyramidal cells 34 . Hence, we would not necessarily detect a change if only a subset of inhibitory synapse types were affected. Moreover, it is not clear whether this specialized synaptic pruning of inhibitory synapses is TREM2-dependent 33 . Overall, the findings of the present study suggest that a decrease in TREM2 function during development, whether due to the Trem2 R47H variant and/or decreased expression, disrupts microglia-mediated synaptic refinement during development, potentially priming hippocampal circuits for heightened vulnerability to hyperactivity and other AD pathology later in life. Specifically, we demonstrate that the Trem2 R47H variant is associated with increased synaptic density in the CA1 and CA3 regions of the hippocampus during early postnatal development, without initially eliciting significant alterations in basal synaptic transmission. These findings suggest that this AD risk variant may influence neuronal connectivity prior to the onset of pathological hallmarks. Limitations of the study The present study has several limitations that warrant consideration. While the presynaptic marker Bassoon is a core component of the active zone at both excitatory and inhibitory synapses 35 , the postsynaptic marker Homer1 is predominantly localized to excitatory synapses, where it functions as a scaffolding protein within the postsynaptic density 36 . Consequently, our synaptic colocalization analysis did not investigate inhibitory synapses, limiting our ability to generalize the observed increase in synaptic density across all synapse types. Furthermore, translational differences between human and murine TREM2 R47H variants must be acknowledged. Specifically, Xiang et al. demonstrated that in this mouse knockin model of the Trem2 R47H variant, aberrant mRNA splicing leads to a decrease of about 60% in Trem2 expression, an effect not observed in human induced pluripotent stem cells-derived microglia or post-mortem brain tissues 10 , 19 . Thus, the effects seen in this study represent not only loss of function due to the R47H mutation but also partial knockdown of TREM2 expression 18 . Moreover, the mice in this study are homozygous for Trem2 R47H, which in principle may make them more relevant to Nasu Hakola disease than AD 3 - 5 . However, the general principle of early effects of microglial dysfunction would apply in both cases and indeed more widely across other forms of neurodegeneration. Future investigations should examine, with ongoing synaptic activity and plasticity as the brain develops, whether these early synaptic changes persist, evolve, or contribute to vulnerability in the context of AD progression, particularly in models that recapitulate amyloid and tau pathology. Resource Availability Lead Contact Requests for additional information, resources, or reagents should be addressed to the lead contact, Frances A. Edwards ( f.a.edwards{at}ucl.ac.uk ). Materials Availability The Trem2 R47H mouse line generated in this study is subject to restrictions imposed by breeder material transfer agreements with The Jackson Laboratory (Bar Harbor, ME, USA). Data and Code Availability All data from this study are available from the lead contact upon request. No original code is reported. Additional information necessary for data reanalysis can be obtained from the lead contact. Authors Contributions Conceptualization: F.A.E. and A.A. Methodology: A.A., D.M.C. and J.W. Investigation: A.A. Visualization: A.A., J.W. and D.M.C. Supervision: F.A.E., D.M.C. and J.W. Writing – original draft: A.A Writing – review & editing: A.A., J.W., D.M.C., and F.A.E. Funding: F.A.E. Declaration of Interests The authors declare no competing interests. STAR⋆Methods Experimental model and study participants details All procedures were performed in accordance with the UK Animals (Scientific Procedures) Act 1986. Trem2 R47H knockin (Jackson Laboratory via MRC Harwell) and C57BL6j WT mice were bred at University College London and used prior to weaning at P18–P22. All mice were housed within individually ventilated cages under a 12:12-hour light–dark cycle at the University College London Biological Services Unit. Food and water were provided ad libitum . Methods details Pups were decapitated and brains bisected in ice-cold ACSF (in mM: 125 NaCl, 1.4 NaH 2 PO 4 , 26 NaHCO3, 2.4 KCl, 20 glucose, 3 MgCl 2 , 0.5 CaCl 2 ; pH 7.4, 310–315 mOsm). One hemisphere was fixed (4% PFA, 24 h, 4 °C) and cryoprotected (30% sucrose in phosphate-buffered saline (PBS)); the other was used for electrophysiology. Electrophysiology Acute transverse hippocampal slices were prepared from P18-22 pups according to Cummings et al. 37 . Briefly, 300 μm-thick slices were prepared in ice-cold dissection ACSF, warmed to 35 °C, equilibrated in ACSF with increasing Ca 2+ (0.5–2 mM) and decreasing Mg 2+ (3–1 mM), and then maintained at room temperature for at least 40 minutes. Whole-cell voltage clamp recordings were performed at V hold = –70 mV from CA1 pyramidal neurons identified using infrared differential interference contrast microscopy (Olympus BX50WI). Pipettes (4–6 MΩ) were filled with internal solution (in mM: 140 CsCl, 2 Mg-ATP, 10 EGTA, 5 HEPES; pH 7.4, ∼290 mOsm). Signals were amplified and low-pass filtered (2 kHz; Multiclamp 700B), digitized (10 kHz; Digidata 1322A) and then acquired within WinEDR (v3.2.7; University of Strathclyde). Series resistance (15–40 MΩ) was monitored throughout. sEPSCs were recorded in ACSF with 6 μM gabazine. mEPSCs were recorded with 1 μM tetrodotoxin added. Events (≥3 pA for ≥2 ms, 10 ms deadtime) were detected using WinEDR and manually confirmed. Event frequency was calculated as 1/inter-event interval; amplitude was peak from baseline. Decay time constants (τ) were obtained by fitting a single exponential from peak to baseline. Paired-pulse ratios were obtained by stimulating Schaffer collaterals (100 μs constant voltage at 25/50/100 ms inter-stimulus intervals. ≥20 repeats were averaged per inter-stimulus interval, with 10 s between each pair of evoked responses) in the presence of gabazine. Pipettes (ACSF-filled) were placed 100–350 μm from the recording site. Only monosynaptic responses (∼4 ms latency) were included in analyses; paired-pulse ratio was calculated as the amplitude of the second response/amplitude of the first response. All recordings and analyses were performed blinded to genotype. Immunohistochemistry Standard protocols were followed to conduct immunohistochemical experiments 38 . Briefly, fixed frozen brain hemispheres embedded in 30% sucrose in PBS were sectioned using a sledge microtome (Leica SM2010R). Sections were cut perpendicular to the hippocampal longitudinal axis at a thickness of 30 μm, immediately transferred to PBS containing 0.02% sodium azide, and stored at 4 °C. Brain sections were permeabilized (0.3% Triton X-100 in PBS), blocked for non-specific binding (3% goat serum in 0.3% Triton X-100 in PBS) and then incubated with primary antibodies (1:500 guinea pig anti-Bassoon, Synaptic Systems, Catalog number 141318; and 1:250 chicken anti-Homer1, Synaptic Systems, Catalog number 160026) at 4 °C for 24 hours, and then with Alexa Fluor-conjugated secondary antibodies (1:500 goat anti-guinea pig AF 594, ThermoFisher, catalog number A-11076; and 1:500 goat anti-chicken AF 647, ThermoFisher, catalog number A-21449) for 2 hours at room temperature. Lastly, sections were mounted onto SuperFrost Plus™ adhesion slides using Fluoromount-G® mounting medium, and then stored at 4 °C. Imaging Two regions of interest were imaged per brain section: a defined area within the stratum radiatum of the CA1 hippocampal subfield, corresponding to Schaffer collateral synapses, and the stratum lucidum of the CA3 region, representing mossy fiber synapses. High-resolution imaging was performed using an Airyscan detector on a Zeiss LSM 880 confocal microscope equipped with a 63× oil-immersion objective lens (NA 1.4). Image acquisition involved averaging four-line scans for the Homer1 and Bassoon channels and a single line scan for the DAPI channel, all captured at 16-bit depth. Z-stacks were acquired with a step size of 0.21 μm across a total depth of 8.01 μm. Laser excitation settings were: 633 nm for Homer1, 561 nm for Bassoon, and 405 nm for DAPI. Photomultiplier tube gain, laser power, and offset settings were optimized during initial calibration and then kept constant across all imaging sessions to ensure consistency. Quantification and statistical analysis Three-dimensional reconstruction of z-stack images for synapse quantification were performed in the Imaris software (Oxford Instruments, Imaris 10.1.1). The “Spots” function was employed to reconstruct Bassoon and Homer1 puncta with a spot size of 0.35 µm, while the “Surfaces” function was used to segment DAPI-stained nuclei. A fixed value threshold was applied uniformly across all sections to ensure consistency in Bassoon and Homer1 spot detection. To exclude nuclear-associated signals, spots located within DAPI-defined surfaces were removed using the in-built exclusion function. Synaptic colocalization was assessed using the “Colocalize Spots” MATLAB XTension, with a colocalization distance threshold set at 0.35 μm. The total number of colocalized Homer1 and Bassoon puncta, representing putative synapses, was quantified. To compute synapse density, the number of colocalized spots outside nuclear (DAPI) volume was divided by the effective tissue volume (i.e., total region of interest volume minus the DAPI-defined volume). Synaptic density values were averaged across three technical replicates to generate a single value per animal. All statistical analyses were conducted using GraphPad Prism 10. Independent t-tests and two-way ANOVA were employed, as specified in the corresponding figure legends. Sample sizes are indicated within the bars of each graph and refer to the number of mice. The threshold for statistical significance was P < 0.05. Acknowledgements This work was supported by grants from Alzheimer’s Research UK and the Cure Alzheimer’s Fund. We thank Haady Hajar for his assistance with mouse colony management in the Edwards Laboratory at University College London. 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