Injection with Toxoplasma gondii protein affects neuron health and survival
A select number of highly divergent microbes ( e . g . , measles virus , Toxoplasma gondii ) naturally cause infections of neurons within the central nervous system ( CNS ) . Most of these viruses cause debilitating disease or death , whereas alphaherpes viruses and the eukaryotic parasite Toxoplasma primarily establish persistent , relatively quiescent neuronal infections in the immunocompetent host ( Dubey , 2009 ) . While neuron-alphaherpes virus interactions have been studied for decades—leading to a mechanistic understanding of herpes virus latency in neurons and the development of novel tools for circuit tracing—our understanding of neuron-Toxoplasma interactions is just beginning ( Ugolini , 1995; Wickersham et al . , 2007 ) . Toxoplasma is a ubiquitous intracellular parasite that can infect a wide range of warm-blooded hosts , including birds , rodents , and humans . In most immunocompetent hosts , Toxoplasma establishes a long-term , asymptomatic infection in specific organs . In humans and rodents , the CNS is a major organ of persistence and is the organ most affected by symptomatic disease in immunocompromised patients ( Remington and Cavanaugh , 1965; Luft and Remington , 1992; Dubey , 2009; Neu et al . , 2015 ) . Using a Cre-based mouse model in which host cells injected with Toxoplasma protein permanently express a green fluorescent protein ( GFP ) , our group showed that Toxoplasma predominantly interacts with neurons , not glia , in vivo ( Koshy et al . , 2012; Cabral et al . , 2016 ) . In addition , we determined that neurons injected with Toxoplasma protein outnumber cysts by over 20-fold ( Koshy et al . , 2012 ) , suggesting that far more neurons interact with Toxoplasma than are persistently infected . Here , we continue to harness this Cre-based model to extend our understanding of the neuron-Toxoplasma interface . We leverage the high numbers of Toxoplasma-injected neurons ( TINs ) and the expression of GFP , which labels the full neuron , to carry out a systematic neuroanatomic mapping of TINs . These studies revealed that TINs are most commonly found within the cortex followed by the striatum and are rarely found in the cerebellum . Within the cortex and the striatum , we used immunofluorescent assays to determine if Toxoplasma preferentially interacts with specific neuron subtypes , finding instead that the lineage of TINs essentially mirrors regional neuron subtype abundance . Finally , we used single neuron patch clamping in ex vivo slices to compare the electrophysiology of striatal TINs to neighboring ‘bystander’ neurons ( neurons within the same slice that were not injected with Toxoplasma proteins ) . These first-of-their-kind studies for any neurotropic microbe showed that TINs—but not bystander neurons—had dramatically altered electrophysiology . At 8 weeks post-infection ( wpi ) , the number of TINs drops by approximately 90% , suggesting this abnormal physiology reflects an unhealthy cellular state . Collectively , these data offer the first insights into which neurons interact with Toxoplasma and how these interactions—either directly or indirectly—alter neuron physiology and survival in vivo .
Prior work in AIDS patients and rodents suggested that Toxoplasma does not evenly distribute across the brain ( Post et al . , 1983; Lang et al . , 1989; Luft and Remington , 1992; Arendt et al . , 1999; Porter and Sande , 1992; Strittmatter et al . , 1992; Berenreiterová et al . , 2011; Evans et al . , 2014; Neu et al . , 2015; Dubey et al . , 2016 ) . These studies have conflicting findings as to where Toxoplasma lesions or cysts ( in rodent studies ) are predominantly found , differences that might be driven by the small numbers of patients or rodents analyzed ( Berenreiterová et al . , 2011; Evans et al . , 2014; Dubey et al . , 2016; Boillat et al . , 2020 ) . Given these conflicting studies , we sought to neuroanatomically map the location of TINs , a population that includes both uninfected , injected neurons and actively infected neurons ( Koshy et al . , 2012 ) . To accomplish this goal , we used our previously published semi-automated method ( Mendez et al . , 2018 ) which utilizes the Allen Institute Mouse Brain Atlas as a reference to quantify and map TINs quickly . This automation made it feasible to sample multiple brain sections per mouse across multiple cohorts of mice infected with either of two genetically distinct Toxoplasma strains ( type II— Prugniaud , type III—CEP ) that express a Toxoplasma:Cre fusion protein and mCherry ( Koshy et al . , 2010; Koshy et al . , 2012; Christian et al . , 2014 ) . For simplicity , from here forward , we will refer to these strains as II-Cre and III-Cre . We quantified TINs in 16–19 brain sections/mouse across 32 regions identified in the Allen Institute Mouse Brain Atlas . Data points were grouped into 12 major regions . For both II-Cre and III-Cre infected mice , the cortex contained the highest average number of TINs , followed by the striatum ( Figure 1A , B; Figure 1—figure supplement 1 ) . In general , III-Cre infected mice showed a higher overall number of TINs compared to II-Cre infected mice ( average of 247 . 5 III-Cre versus 216 . 7 II-Cre ) . The increase in absolute numbers in III-Cre infected mice was most pronounced in the cortex ( Figure 1A , B ) . Most other brain regions , including the cerebellum , contained relatively few TINs ( Figure 1A , B ) . As the cortex and striatum encompass a large proportion of the brain , the high number of TINs in these areas might be driven by relative size . To test this possibility , we produced an enrichment index using the following equation: RtinTtinRsize , where Rtin is the TIN count for a given region ( e . g . , cortex ) , Ttin is the total TIN count from an individual mouse , and Rsize is the total area percentage of a specific region ( e . g . , % of brain that is cortex ) ( see Supplementary file 1 for area percentage for each brain region ) . Our method of sectioning does not allow us to collect the smaller sections of the cerebellum , therefore we only included the cerebellar areas from sections 10-16 of the Allen atlas for normalization ( consistent with the regions of the cerebellum which were analyzed ) . To align our sections with the Allen Institute sections , we also excluded the olfactory bulb ( Mendez et al . , 2018 ) . Even with these exclusions , 85% of the original Allen atlas area was used . If the number of TINs in a region is proportional to the size of that region , the normalization index will approximate 1 , while regions enriched for TINs will have indices > 1 , and regions relatively devoid of TINs will have indices < 1 . Using this normalization , we determined that the cortex was the only region which was significantly enriched for TINs in both II-Cre and III-Cre infected mice ( index score 1 . 4 ± 0 . 1 and 2 . 2 ± 0 . 2 , respectively ) ( Figure 1C , D ) . For both groups of infected mice , within the cortex , the somatosensory , motor , and visual cortices had the highest enrichment score ( Figure 1—figure supplement 2 ) . Finally , two regions—the cerebellum and the white matter tracts—showed a significant lack of enrichment in both II-Cre and III-Cre infected mice ( Figure 1C , D ) . The lack of TINs in white matter tracts is expected given that these tracts are primarily made up of neuron axons and thus have a paucity of neuron nuclei , the cellular component by which neurons are anatomically mapped . Supplementary file 2 has the full list of enrichment scores and statistical analysis for the 12 listed regions . In summary , these data show that , irrespective of infecting Toxoplasma strain , TINs are commonly found in the cortex followed by the striatum and relatively rarely found in the cerebellum . While TINs localization in the striatum is consistent with its size , the enrichment of TINs in the cortex and lack of TINs in the cerebellum are not accounted for by the relative size of these regions . As the cortex and the striatum had the highest number of TINs , we decided to use these areas to determine if Toxoplasma targeted a specific neuron population . Neurons can be classified in many ways , including by neurotransmitter expression , physiology , and morphology . The simplest way is to classify neurons is as inhibitory or excitatory . Given that prior work suggested that Toxoplasma infection might specifically affect inhibitory neurons ( Brooks et al . , 2015 ) , we first examined how often TINs co-localized with two common markers for inhibitory interneurons , calbindin and parvalbumin . We chose these makers rather than the pan-inhibitory marker glutamate decarboxylase ( GAD ) [GAD+ interneurons make up approximately 15–20% of all cortical neurons ( Gentet et al . , 2000; Lodato and Arlotta , 2015 ) ] as GAD staining is altered in Toxoplasma-infected brain ( Brooks et al . , 2015; Carrillo et al . , 2020 ) . Fortunately , parvalbumin labels 40–50% of GAD+ interneurons and calbindin labels approximately 30% of GAD+ interneurons ( Hof et al . , 1999; Keller et al . , 2019 ) . To quantify how often TINs were calbindin or parvalbumin interneurons , we labeled brain sections with antibodies against either calbindin or parvalbumin , and with antibodies against NeuN , a pan-neuronal marker . These labeled sections were then imaged with confocal microscopy to identify TINs that co-stained with NeuN and either calbindin or parvalbumin . Within the cortex , irrespective of infecting strain , approximately 5% ( 4 . 8 ± 2 . 0% II-Cre , 3 . 9 ± 0 . 6% III-Cre ) of TINs were calbindin+ ( Figure 2A , B ) , while none were parvalbumin+ ( Figure 2C , D ) . In the striatum , we observed that approximately 4% of II-Cre TINs were calbindin+ ( 4 . 4 ± 2 . 0% ) and approximately 9% of III-Cre TINs were calbindin+ ( 9 . 4 ± 3 . 4% ) ( Figure 2E , F ) . As in the cortex , no TINs co-localized with parvalbumin+ neurons in the striatum ( N = 6–65 TINs/mouse , four mice for II-Cre; N = 30 and 84 TINs/mouse , two mice for III-Cre ) . These data suggest that Toxoplasma rarely interacts with or injects cortical or striatal interneurons , though we cannot exclude the possibility that Toxoplasma-injected interneurons die prior to the expression of GFP or that parasites primarily interact with interneurons that do not express parvalbumin or calbindin . Either scenario would limit our ability to detect these Toxoplasma-interneuron interactions . Within interneurons , Toxoplasma more commonly injects calbindin+ interneurons , not parvalbumin+ interneurons , an unexpected finding given the higher prevalence of parvalbumin+ interneurons . As only a small number of TINs co-localized with markers for inhibitory neurons , we next sought to determine if TINs co-localized with a marker for excitatory cortical neurons . For these studies , we selected the transcription factor FoxP2 . FoxP2 is primarily expressed by glutamatergic/excitatory neurons within layer 6 of the mouse cortex , though some FoxP2+ excitatory neurons are also found in cortical layers IV and V ( Arlotta et al . , 2005; Lodato and Arlotta , 2015 ) . FoxP2 expression also occurs in medium spiny neurons ( MSNs ) in the striatum , allowing us to use a single marker for both cortical and striatal studies ( Ferland et al . , 2003; Lai et al . , 2003 ) . We quantified how often TINs were FoxP2+ by labeling and imaging brain sections as above except now using antibodies directed against FoxP2 . Within the cortex , irrespective of infecting Toxoplasma strain , approximately 58% ( 58 . 0 ± 3 . 8% II-Cre , 58 . 3 ± 2 . 3% III-Cre ) of the analyzed TINs showed FoxP2 co-localization ( Figure 3A , B ) . In the striatum , we found that approximately 68% ( 68 . 1 ± 7 . 2% ) of analyzed II-Cre TINs and approximately 83% ( 83 ± 4 . 1% ) of analyzed III-Cre TINs showed FoxP2 co-localization ( Figure 3C , D ) . This high rate of co-localization between striatal TINs and FoxP2 suggested that striatal TINs were MSNs , which would be expected given that 90–95% of neurons in the striatum are MSNs ( Ouimet et al . , 1984; Ouimet and Greengard , 1990 ) . As the identification of striatal TINs as MSNs had important implications for pursuing electrophysiology studies , we sought to further confirm that striatal TINs were MSNs by determining the rate of co-localization between striatal TINs and DARPP32 , another marker for MSNs ( Ouimet et al . , 1984; Ouimet and Greengard , 1990 ) . We found that approximately 20% ( 19 . 9 ± 5 . 7% ) of II-Cre striatal TINs and approximately 25% ( 23 . 6 ± 4 . 8% ) of III-Cre striatal TINs co-localized with DARPP32+ staining ( Figure 3E , F ) . Given the relatively low percentage of TINs co-localizing with DARPP32 and prior work suggesting Toxoplasma infection alters protein localization and expression ( Cekanaviciute et al . , 2014; Brooks et al . , 2015; David et al . , 2016 ) , we sought to determine how striatal FoxP2 and DARPP32 staining differed between uninfected and infected mice , and if this staining differed by proximity to TINs . To determine the abundance of FoxP2 and DARPP32 neurons near TINs , we re-analyzed the images used for Figure 3 , this time quantifying the number of GFP- neurons ( NeuN+ cells ) that co-localized with FoxP2 or DARPP32 . To determine the abundance of FoxP2 and DARPP32 neurons regardless of proximity to TINs , we generated new , randomly distributed images from a subset of stained sections used for Figure 3 and quantified the co-localization between GFP- neurons and FoxP2 or DARPP32 . We found that the average number of FoxP2+ neurons in each image did not differ between images from either uninfected or infected mice , regardless of infecting Toxoplasma strain ( Figure 3—figure supplement 1A ) ; nor did the abundance vary in proximity to TINs ( Figure 3—figure supplement 1B ) . The average number of DARPP32+ neurons in each image also did not differ between random images from uninfected or infected mice ( Figure 3—figure supplement 1C ) . Conversely , in images taken in proximity to TINs , the number of DARPP32+ neurons showed a statistically significant decrease ( Figure 3—figure supplement 1D ) . In addition , we noted that , on average , more striatal neurons were FoxP2+ than DARPP32+ , even in uninfected mice ( Figure 3—figure supplement 1A , C ) . To further test the possibility that FoxP2 identifies more striatal neurons than DARPP32 , we stained tissues sections for both FoxP2 and DARPP32 and compared the co-localization between these proteins . Consistent with the findings in the single stains , only approximately 70% of FoxP2+ neurons stained for DARPP32 ( 69 . 6 ± 0 . 3% saline , 68 . 6 ± 0 . 6% II-Cre , 73 . 0 ± 4 . 0% III-Cre ) ( Figure 3—figure supplement 2A ) , while 99% ( 99 . 7 ± 0 . 0% saline , 99 . 8 ± 0 . 0% II-Cre , 99 . 8 ± 0 . 1% III-Cre ) of DARPP32+ neurons stained for FoxP2 ( Figure 3—figure supplement 2B ) . These data suggest that striatal TINs are likely MSNs as they co-localize with markers of MSNs , especially FoxP2 . The higher co-localization of TINs with FoxP2 versus DARPP32 is likely driven by the higher number of FoxP2+ neurons within the striatum as well as the sensitivity of DARPP32 expression/staining to disruption by infection or inflammation . Given that FoxP2 stains more striatal neurons than DARPP32 , even in uninfected mice , FoxP2 may be a more sensitive marker for striatal MSNs than DARPP32 . The identification of striatal TINs as co-localizing with markers for MSNs offered us an unusual opportunity . MSNs make up 90–95% of dorsal striatal neurons ( Mensah and Deadwyler , 1974; Graveland and DiFiglia , 1985; Gerfen , 1992 ) and are highly characterized , including at the level of single cell electrophysiology in situ ( i . e . , patch clamping in ex vivo slices ) . Targeting such a well-characterized , high-frequency population would allow us to directly compare the electrophysiology of TINs and bystander neurons—striatal neurons in proximity to a TIN but not injected with Toxoplasma protein—as both groups of neurons would likely be MSNs . We reasoned that comparing the physiology of TINs and bystanders would allow us to determine the role of general inflammation in driving any changes we observed in TINs ( i . e . , if TINs and bystanders showed similar physiologic changes , then these changes are likely driven by the general inflammatory response to Toxoplasma , not from direct manipulation of TINs by Toxoplasma effector proteins ) . For these studies , we chose to record from III-Cre infected mice because the type III strain produces an higher frequency of TINs than the type II strain ( Figure 1A ) , and because III-Cre TINs show higher rates of co-localization with the MSN marker FoxP2 ( Figure 3F ) . To ensure our capability to patch-clamp onto MSNs in ex vivo slices , we obtained thick brain slices from uninfected mice and performed patch clamping using a standard protocol ( Nisenbaum et al . , 1994; Suter et al . , 1999; Haubensak et al . , 2010; Wang et al . , 2019 ) . As expected , in uninfected mice , dorsal striatum neurons showed classical MSN electrophysiologic properties such as a hyperpolarized resting membrane potential of approximately −80 mV and a delayed first action potential ( AP ) ( Figure 4A ) . In addition , when we filled a neuron showing the above characteristics and stained the section for DARPP32 , we found the filled neuron co-localized with DARPP32 labeling ( Figure 4B–D ) . In infected mice , we electrophysiologically interrogated bystander MSNs and TINs , with bystander MSNs being approximately 30–100 µm from a TIN . Bystander MSNs had a resting membrane potential of −69 . 0 ± 1 . 2 mV , which is mildly depolarized compared to MSNs in uninfected mice ( Figure 5A , Figure 5—figure supplement 1 ) . Consistent with a depolarized resting membrane potential , bystander MSNs required fewer steps of injected current to reach the threshold to fire the first AP compared to MSNs in uninfected mice ( 12 ± 0 . 3 steps for bystanders and 13 ± 0 . 4 for MSNs in uninfected mice ) ( Figure 5—figure supplement 2 ) . Besides these two properties , the following electrophysiologic properties were equivalent when comparing bystander MSNs to MSNs from uninfected mice: AP threshold , after hyperpolarization peak , AP amplitude , delay to first AP , and input resistance ( Figure 5B–G ) . These results indicate that though there is a mild change in the resting membrane potential of bystander MSNs during infection , the infected/inflammatory microenvironment at this time point causes relatively little change to neurons that have not directly interacted with Toxoplasma . We next evaluated TINs by following the same procedure as when we recorded from uninfected and bystander MSNs , except that we used the expression of GFP to identify the TIN ( Figure 6—figure supplement 1 ) . Unexpectedly , we were unable to properly patch onto most striatal TINs ( i . e . , we were unable to form a gigaseal ) . We were able to record from a total of 10 TINs , most of which had highly depolarized resting membrane potentials ( −49 . 1 ± 4 . 5 mV ) compared to either bystander or uninfected MSNs ( Figure 6A ) . None of these TINs generated APs in response to the standard protocol , possibly because this highly depolarized membrane potential is too close to the threshold for firing an AP ( ~45 mV ) in MSNs ( Figure 5B ) . Several possibilities could most easily explain this highly depolarized membraned potential: ( i ) the patched GFP+ cells were not neurons , ( ii ) the patched cells were dead neurons , or ( iii ) the patched cells were unhealthy neurons . To distinguish between these possibilities , when possible , we hyperpolarized the patched TINs , followed by attempts to generate APs . For five TINs , after we applied between −20 and −180 pA of negative current , membrane potentials of −55 to −66 mV were observed . With this ‘normalization’ of the resting membrane potential , we found that TINs could generate APs ( Figure 6B ) , definitively showing that they were neurons . Finally , to ensure the depolarized resting membrane potential of TINs was not due to fluorescent protein expression in the setting of inflammation , we recorded from YFP+ neurons in infected mice that constitutively express YFP in a subset of cortical neurons ( Feng et al . , 2000 ) . In these mice , we found a range of resting membrane potentials that is typical of cortical neurons ( Pangratz-Fuehrer and Hestrin , 2011; Oswald et al . , 2013 ) and distinct from the physiologic abnormalities observed in TINs ( Figure 6—figure supplement 2 ) . Given that such findings are consistent with the electrophysiology of sick or dying neurons ( Lipton , 1999 ) , we compared the total number of TINs in brain sections from mice infected with III-Cre parasites for 3 or 8 weeks . Consistent with the possibility that the electrophysiology reflects that TINs are sick/dying , brain sections from 8 wpi mice showed an approximately 90% reduction in the number of TINs compared to brain sections from three wpi mice ( Figure 7 ) . Collectively , our data suggest that neurons that directly interact with Toxoplasma have highly aberrant electrophysiology and most will die over a relatively short period . On the other hand , bystander neurons , which are in a similar inflammatory environment , show relatively minor changes in terms of electrophysiology . Together , these data suggest that neuron pathology is extremely targeted , even in a highly inflammatory setting .
In this study , we leveraged the Toxoplasma-Cre system to extend our understanding of neuron-Toxoplasma interactions . Across multiple cohorts of mice and using two genetically distinct Toxoplasma strains , our data show that TINs consistently localize to the cortex and striatum . For the cortex , the number of TINs is higher than would be expected even after accounting for the size of the cortex . Within the cortex and striatum , we found little evidence of Toxoplasma targeting a specific neuron subtype , as TINs co-localize with markers for highly abundant neurons in a region ( e . g . , MSNs in the striatum ) . Interestingly , when we used whole cell patch clamping in ex vivo slices to assess both striatal TINs and bystander MSNs , we found discrepant electrophysiology . Bystander MSNs showed mild changes in resting membrane potential and no changes in other passive firing properties , while TINs had highly depolarized resting membrane potentials and did not fire APs unless artificially hyperpolarized . These data suggest that TINs , but not bystanders , are unhealthy and potentially dying , a possibility confirmed by a drastic decrease in the number of TINs by 8 wpi . Together , these data offer an unprecedented look into the neuron-Toxoplasma interface , raising new questions about why the cortex is particularly vulnerable to Toxoplasma infection and the evolutionary advantage of the death of neurons injected with Toxoplasma protein . One of the major advances of this work is how we mapped neuron-Toxoplasma interactions . Most previous studies have utilized a single Toxoplasma strain , relatively few animals ( or more animals but relatively few sections/animal ) , and were limited to measuring cyst location . On the other hand , we mapped TINs , which outnumber cysts ( Koshy et al . , 2012 ) and show uniform distribution of GFP , allowing us to map neuron cell bodies/somas which is the traditional method for identifying neuron anatomic location . Furthermore , as our semi-automated method is relatively quick ( Mendez et al . , 2018 ) , in total , we mapped a total of over 5000 TINs/Toxoplasma strain , using multiple cohorts of mice and >15 sections/mouse . These robust methods offer confidence that our findings are not unduly influenced by a single outlier mouse , a cohort effect , or insufficient sampling for a process that is highly heterogenous . Our findings are consistent with several aspects of the two most extensive cyst mapping studies ( Berenreiterová et al . , 2011; Boillat et al . , 2020 ) . All three studies found: ( i ) high levels of inter-mouse variability; ( ii ) identified the cortex—and especially the motor , somatosensory , visual cortical regions—as showing enrichment for Toxoplasma presence ( TINs or cysts ) even when accounting for region size; and ( iii ) determined that the cerebellum shows relatively little Toxoplasma presence , especially when accounting for size . These consistencies suggest that regional cyst burden is driven by differences in the number of neuron-parasite interactions , not by differences in regional innate immune responses as has been suggested for West Nile virus ( WNV ) infection ( Cho et al . , 2013 ) . In addition , these regional differences are consistent with the largest studies in AIDS patients with toxoplasmic encephalitis , in which more lesions are identified in the cortex and the basal ganglia compared to the cerebellum ( Porter and Sande , 1992; Arendt et al . , 1999 ) , highlighting the relevance of the mouse model to human disease . Currently , we do not know what drives these regional differences in tropism , though one possibility is differences in the transmissibility of Toxoplasma across the vasculature that supplies the cortex and striatum/basal ganglia versus the cerebellum ( anterior versus posterior circulation , respectively ) , a possibility also suggested for WNV ( Daniels et al . , 2017 ) . A second advance in our study is the ability to identify the type of neuron injected by Toxoplasma . The finding that TINs are comprised of common neuron subtypes in the deep cortex and striatum ( e . g . , FoxP2+ neurons ) suggests that Toxoplasma injects ( and presumptively infects ) in proportion to the neuron subtypes in a given region . One exception to this rule is the lack of TINs that also express parvalbumin . Parvalbumin-expressing neurons make up approximately 40–50% of the cortical GABAergic ( inhibitory ) neurons and are found in all deeper layers of the cortex , while calbindin-expressing neurons are less frequent , especially in layers IV-VI of the cortex ( Hof et al . , 1999; Tremblay et al . , 2016 ) . One possibility for the lack of parvalbumin+ TINs is a decrease or change in parvalbumin expression/location as seen for DARPP-32 ( Figure 3—figure supplement 1 ) or GAD-67 ( Brooks et al . , 2015 ) . Another more exciting possibility is that parvalbumin+ neurons have fewer interactions with Toxoplasma because they have a high concentration of perineural nets—extracellular matrix that surrounds different parts of the neuron ( Baker et al . , 2017 ) —blocking Toxoplasma’s ability to contact parvalbumin+ neurons . Another possibility is that parvalbumin+ neurons are highly sensitive to injection with Toxoplasma protein and rapidly die before expressing GFP and/or die prior to the time point at which we performed our studies . Future studies will define the factors that determine which neurons interact with parasites . A final advance in this work is the electrophysiology comparing TINs and bystander neurons . To the best of our knowledge , these studies are the first in any neurotropic infection to compare the in situ electrophysiology between microbe-interacted ( or infected ) neurons versus those neurons in close proximity but without direct microbial interactions . The utility of comparing TINs and bystanders is to address what physiologic effects are driven by the general neuroinflammatory response versus those changes driven by neuron-parasite interactions . Given the relatively common understanding that microglia will strip synapses off infected and uninfected neurons in the inflamed brain ( Vasek et al . , 2016; Di Liberto et al . , 2018 ) , we found it remarkable that bystander MSNs physiology showed only minor changes compared to MSNs from uninfected mice while TINs showed highly abnormal electrophysiology ( Figure 6 ) . One possible explanation for the minimal changes seen in bystanders is that these neurons undergo compensatory mechanisms that allow them to avoid drastically changing , as a dramatic response would be detrimental to the local circuitry . Another possibility is that the effect of the neuroinflammatory response is confined to a small area near TINs , leaving these more distant bystanders relatively spared . Either explanation would preserve circuit function at a global level and allow for a relatively quick reversal of these physiologic changes once the inflammatory response subsides . Of note , while the physiology of the bystanders is relatively normal compared to TINs , this depolarized resting membrane potential is still abnormal and the decrease in required input current ( Figure 6—figure supplement 2 ) suggests these neurons are mildly hyperexcitable . If similar changes also occur in other bystander neurons ( e . g . , cortical neurons ) and in other CNS infections , they might explain why patients with active brain infections have an increased likelihood of seizing ( Luft and Remington , 1992; Neu et al . , 2015 ) . These mild abnormalities might also summate across many bystander neurons to cause circuit level changes . While we did not test this possibility , such changes in the far more numerous bystanders—especially if the changes are reversible—could explain both the heterogeneity of behavioral changes seen in infected animals ( Worth et al . , 2014 ) and the relative lack of behavioral abnormalities observed in Toxoplasma-infected mice that have cleared both the infection and the immune response ( McGovern et al . , 2020 ) . TINs—which include actively infected neurons and uninfected , injected neurons ( Figure 5—figure supplement 1 ) —show highly abnormal electrophysiology , only firing APs after artificial hyperpolarization . While we cannot distinguish between whether this abnormal physiology leads to neuron death or the sick state of these neurons drives the abnormal physiology ( we favor the latter possibility ) , the discrepancy between the physiology of TINs and bystanders suggests that the root cause is specific to neurons injected with Toxoplasma protein . Such restricted changes could arise from direct manipulation of TINs by the injected parasite proteins or selective microglial/macrophage recognition and phagocytosis of TINs , consistent with what was observed in lymphocytic choriomeningitis virus infection ( Di Liberto et al . , 2018 ) . Another possibility , as noted above , is that the immune response is tightly regulated and confined , such that the destruction leveled by this response is limited to a very small area around TINs . Such a possibility is consistent with prior work showing that partial abrogation of astrocytic TGF-β signaling led to de-regulation of the CNS immune response to Toxoplasma and higher levels of neuronal loss without changing the CNS parasite burden ( Cekanaviciute et al . , 2014 ) . Future studies will determine what factors drive this abnormal physiology . Finally , our finding of an approximately 90% decrease in TINs by 8 wpi suggests that we need to re-evaluate our understanding of how and why microbes persist in or even infect neurons . Microbes with the capability to infect multiple cell types are commonly found in neurons , not astrocytes ( e . g . , Toxoplasma , WNV , measles virus ) . This neuronal preference has been explained by the idea that—given the long-lived , relatively non-regenerative nature of neurons—neuron survival must be ensured at almost any cost . Thus , rather than initiating an immune response that might result in neuron death , neurons survive infection through non-cytolytic clearance of microbes ( Binder and Griffin , 2001; Patterson et al . , 2002; Samuel and Diamond , 2005; Miller et al . , 2016 ) and/or pushing the infecting microbes to enter a latent state ( Speck and Simmons , 1991; Ferguson and Hutchison , 1987; Tanaka et al . , 2016 ) . The death of TINs directly challenges this model , suggesting that the battle to persist in neurons is far more nuanced than previously recognized . For example , given that >90% of TINs are expected to be uninfected ( Koshy et al . , 2012 ) , we would predict that many uninfected neurons are dying . What is the advantage of uninfected , parasite-manipulated neurons dying ? One possibility is that these TINs are so abnormal ( from either direct manipulation by parasites or because of neuron-immune cell interactions ) that global brain function is better preserved with the removal of these neurons . Such a possibility is consistent with the altered gene expression identified in neurons that cleared an attenuated rabies virus infection ( Gomme et al . , 2012 ) . Another possibility is that the death of TINs—both infected and uninfected—is essential for parasite control and perhaps the death of uninfected TINs is simply collateral damage . Or the death of uninfected TINs might divert immune attention from the rare , persistently infected neuron . Regardless of whether this death benefits the host or microbe ( or both ) , based upon the data presented here and previously , we offer an alternative model for neuron-tropic microbes: neurons mount appropriate immune responses to both cytokines and infections such that persistent infection is a rare outcome , rather than a universal one . Limited data in a murine model of measles virus infection is consistent with this possibility ( Miller et al . , 2019 ) , though future studies using other neurotropic pathogens will be required to definitively determine how universal this model is .
Parasites were maintained via serial passage through human foreskin fibroblasts using DMEM supplemented with 2 mM glutagro , 10% fetal bovine serum , and 100 IU/ml penicillin/100 μg/ml streptomycin . The type II ( Prugniaud ) and type III ( CEP ) strains used have been engineered to express mCherry and Cre recombinase and have been previously described ( Koshy et al . , 2012; Christian et al . , 2014 ) . Most mice used in these studies are Cre-reporter mice that express GFP only after the cells have undergone Cre-mediated recombination ( Madisen et al . , 2010 ) . For Figure 6—figure supplement 2 ( recordings of YFP+ cortical neurons ) , thy1-YFP-H mice were used ( Feng et al . , 2000 ) . Mice were originally purchased from Jackson Laboratories ( stock # 007906 for Cre-reporter mice , stock # 003782 for thy1-YFP mice ) and bred in the University of Arizona Animal Care Facilities . Mice were infected at 2–3 months of age via intraperitoneal injection with freshly syringe-released parasites for Figures 1 , 2 , 3 and 7 , and related figure supplements . Mice were inoculated with 10 , 000 freshly syringe-released parasites in 200 μl of USP-grade PBS for both II-Cre and III-Cre strains . To increase the reliability of electrophysiology studies ( Figures 5 and 6 , and related Figure 6—figure supplements 1 and 2 ) , mice were infected at 5–6 weeks of age with III-Cre parasites only . For the localization studies , at 3 wpi animals were sedated with a ketamine/xylazine cocktail , intracardially perfused with saline followed by 4% paraformaldehyde ( PFA ) , after which brains were harvested . Sections were then prepared as previously described ( Mendez et al . , 2018 ) . In brief , left and right brain hemispheres were isolated , drop-fixed in 4% PFA followed by cryopreservation in 30% sucrose . Forty-micron-thick sagittal sections were generated using a freezing sliding microtome ( Microm HM 430 ) . Sections were then stored as free-floating sections in cryoprotective media ( 0 . 05 M sodium phosphate buffer containing 30% glycerol and 30% ethylene glycol ) until labeling procedure was to be done . Sections for localization studies ( Figure 1 , Figure 1—figure supplements 1 and 2 ) were sampled every 200 μm , while for co-localization studies ( Figures 2 and 3 , Figure 3—figure supplements 1 and 2 ) , sections were sampled every 400 μm . To ensure adhesion of tissue onto slides for localization studies , tissue was air-dried onto slides overnight then heated on a slide warmer for 40 min at 34°C . Then tissue was dehydrated using increasing then decreasing concentrations of 50% , 75% , 95% , and 100% ethanol . Slides were washed with TBS , peroxidase inactivated ( 3%H2O2/10% methanol ) , permeabilized ( 0 . 6% Triton X-100 ) , blocked ( 1 . 5% BSA and 1 . 5% goat serum ) , and incubated with rabbit anti-ZsGreen ( Clontech , Cat . No . 632474 , 1:10 , 000 ) for 15–18 hr at room temperature . Next , slides were incubated in goat anti-rabbit polyclonal biotinylated conjugated antibody ( Vector Labs , Cat . No . BA-1000 , 1:500 ) for 2 hr , followed by incubation with avidin-biotin complex kit ( Thermo Fischer Scientific , 32020 ) for 2 hr and visualization with a 3 , 3’-diaminobenzidine kit ( Vectastain , Vector Labs , SK-4100 ) . Sections were then counterstained with cresyl violet for Nissl labeling . For co-localization studies , sections were labeled free-floating , 8–10 sections/mouse . Sections were washed with PBS , permeabilized with 0 . 3% Triton-X , then permeabilized and blocked with 0 . 3% Triton-X and 5% goat serum ( Jackson Immuno ) . For primary antibodies , the following were incubated for 15–18 hr at 4°C in PBS with 0 . 3% Triton-X and 5% goat serum: mouse biotin conjugated anti-NeuN B-clone A60 ( Millipore , MAB377 , 1:500 ) , rabbit anti-calbindin ( Sigma-Aldrich , C2724 , 1:500 ) , rabbit anti-parvalbumin ( Abcam , ab11427 , 1:1000 ) , rabbit anti-FoxP2 ( Abcam , ab16046 , 1:1000 ) , rabbit anti-DARPP32 ( Abcam , ab40801 , 1:500 ) . Following incubation in the appropriate antibody , sections were then incubated for 2 hr in the following secondaries: goat anti-rabbit IgG Alexa Fluor 568 ( Invitrogen , A11011 , 1:500 ) , Cy5 streptavidin ( Invitrogen , SA1011 , 1:500 ) , and all sections were counterstained with 4’ , 6-diamidino-2-phenylindole ( DAPI ) ( Thermo Fisher Scientific , 1:1000 ) . For filled MSNs , sections were processed as discussed below . For staining , sections with filled MSNs were permeabilized with 0 . 6% Triton-X in PBS , then incubated with DARPP32 antibody in 5% goat serum , 0 . 6% Triton-X in PBS for 48 hr at 4°C . Sections were washed then incubated with Cy5 streptavidin ( 1:500 ) and goat anti-rabbit Alexa Fluor 568 ( 1:500 ) for 24 hr at 4°C . Mouse brain slice electrophysiology recording was performed as described ( Suter et al . , 1999; Nisenbaum et al . , 1994; Dorris et al . , 2014; Willett et al . , 2016; Haubensak et al . , 2010 ) . Mice were sacrificed with CO2 , after which the descending aorta was clamped and the mice were intracardially perfused with ice-cold oxygenated artificial cerebral spinal fluid ( ACSF ) containing 126 mM NaCl , 1 . 6 mM KCl , 1 . 2 mM NaH2PO4 , 1 . 2 mM MgCl2 , 2 . 4 mM CaCl2 , 18 mM NaHCO3 , 11 mM glucose . After the olfactory bulbs and the cerebellum were removed , the brain was transferred to a vibratome stage that contained ice-cold ACSF oxygenated with carbogen ( 95% O2 balanced with CO2 ) ; 200 μm coronal sections were then generated with the vibratome ( Leica , VT1000S ) . Brain slices were immediately transferred to oxygenated NMDG-HEPES recovery solution ( 93 mM NMDG , 2 . 5 mM KCl , 1 . 2 mM NaH2PO4 , 30 mM NaHCO3 , 20 mM HEPES , 25 mM glucose , 5 mM sodium ascorbate , 2 mM thiourea , 3 mM sodium pyruvate , 10 mM MgSO4 , 0 . 5 mM CaCl2 , 300–310 mOsm , titrated with 10 N HCl to adjust pH to 7 . 3–7 . 4 ) and were allowed to recover for 15 min at 32–34°C . After the recovery portion , brain slices were then transferred to room temperature oxygenated ACSF for 1 hr . After the rest period , recordings were performed in a rig equipped with a MultiClamp 700B , a Digidata 1550A1 ( Molecular Devices ) , and a fluorescent microscope ( Olympus BX51 ) that was used to visualize sections and identify TINs . Patch pipettes were pulled with P-97 Sutter micropipette puller to achieve a resistance of 8–16 MΩ , after which they were filled with an intracellular solution ( 135 mM potassium gluconate , 5 mM EGTA , 0 . 5 mM CaCl2 , 2 mM MgCl2 , 10 mM HEPES , 2 mM MgATP , and 0 . 1 mM GTP , pH 7 . 3–7 . 4 , 290–300 mOsm ) . Recording data were sampled at 10 kHz , filtered at 3 kHz , and analyzed ( Molecular Devices , pCLAMP , 10 . 7 ) . For bystander neurons and neurons in uninfected mice , the classification as an MSN was based on the firing pattern in response to the current injections as described previously ( Schmidt and Perkel , 1998; Farries and Perkel , 2000; Luo et al . , 2001; Ding and Perkel , 2002; Ding et al . , 2003 , Willett et al . , 2016 ) . For filling experiments , once data from a 200 μm section were recorded , the recorded neurons were filled with 0 . 6% neurobiotin ( Vector Labs , SP1120 ) . Filled sections were then stored in 4% PFA for 24 hr followed by cryopreservation in cryoprotective media until stained as described above . Slides for localization data were imaged on a Leica DMI6000 with a motorized stage , using Leica Application Suite X ( LAS X ) at 10× magnification . Base background subtraction and white balance was maintained throughout individual cohorts . Image stitching was done automatically through LAS X with a 10% overlap and images stored as Leica image file format ( lif ) . Lif images were then processed on a custom MATLAB code for image transformation onto the Allen Institute reference atlas , followed by semi-automated quantification , as described in Mendez et al . , 2018 . Animals were excluded due to having the same or fewer TINs compared to saline-injected animals ( ~15 TINs ) and via ROUT outlier tests . Figure 1—figure supplement 1 has all mice included and includes 100% of the Allen atlas for the enrichment index . For co-localization studies , 8–10 sections/mouse were imaged . Images were captured on a Zeiss LSM880 inverted confocal microscope ( Marley Microscopy Core , UA ) . Images were captured with a 20× objective , with 1× PMT zoom , all settings were the same across specific staining runs . Autofocus was used to capture a z-stack that would capture from edge-to-edge of the section . In Figure 1 , Figure 1—figure supplement 2 , data were analyzed with a one-sample t-test to compare the enrichment score to a value of 1 , which would indicate a random distribution of TINs in a given region of the brain . For Figures 2–3 and associated figure supplements , the data were analyzed with an unpaired two-tailed t-test to test for differences between II-Cre and III-Cre infected groups . For Figure 5 and Figure 7 , the data were analyzed with a Mann-Whitney U-test . Figure 5—figure supplement 2 was analyzed with an unpaired two-tailed t-test . Figure 6 was analyzed with a one-way ANOVA with Tukey’s post-test . All statistics were done via Prism statistical software ( GraphPad v9 . 1 . 0 ) .
Toxoplasma gondii is an intracellular parasite that causes a long-term latent infection of neurons . Using a custom MATLAB-based mapping program in combination with a mouse model that allows us to permanently mark neurons injected with parasite proteins , we found that Toxoplasma-injected neurons ( TINs ) are heterogeneously distributed in the brain , primarily localizing to the cortex followed by the striatum . In addition , we determined that cortical TINs are commonly ( >50% ) excitatory neurons ( FoxP2+ ) and that striatal TINs are often ( >65% ) medium spiny neurons ( MSNs ) ( FoxP2+ ) . By performing single neuron patch clamping on striatal TINs and neighboring uninfected MSNs , we discovered that TINs have highly aberrant electrophysiology . As approximately 90% of TINs will die by 8 weeks post-infection , this abnormal physiology suggests that injection with Toxoplasma protein—either directly or indirectly—affects neuronal health and survival . Collectively , these data offer the first insights into which neurons interact with Toxoplasma and how these interactions alter neuron physiology in vivo .
Toxoplasma gondii is an intracellular parasite that infects the brain . Whereas most microbial infections of the brain result in severe illness or death , Toxoplasma gondii infections are usually asymptomatic . This is because the parasite has evolved the ability to exist within the brain by dampening the immune response . The parasite can therefore asymptomatically co-exist with its host for years – or even an entire lifetime . The strategy has proved so successful that up to one third of the world’s population is now thought to be infected with Toxoplasma gondii . While this persistence tends not to be a problem for most healthy individuals , dormant Toxoplasma gondii parasites can reactivate in individuals whose immune systems fail . This can result in life-threatening neurological disease . In pregnant women , Toxoplasma gondii parasites can also cross the placenta , which can trigger miscarriage or cause harmful disease in the newborn . To develop treatments for these cases of symptomatic disease , we need to understand how the parasite hides from the immune system in asymptomatic individuals . Mendez et al . have therefore leveraged a mouse model in which neurons injected with Toxoplasma gondii proteins ( Toxoplasma-injected neurons , or ‘TINs’ ) produce a green fluorescent protein . This enables the infected cells to be viewed under a microscope . Examining the mouse brains revealed that most TINs were located in two specific regions: the cortex and the striatum . The cortex is the brain’s outer layer of tissue . The striatum is a structure deep within the brain that helps regulate movement and responses to rewards . Both the cortex and the striatum contain different types of neurons . The results revealed that the proteins from the parasite were spread roughly equally among the various cell types , rather than targeting a specific subtype of neuron . Neurons close to TINs had slightly abnormal electrical activity , whereas the TINs themselves had highly abnormal activity . By eight weeks post-infection , however , the number of TINS had fallen by around 90% . This suggests that many neurons containing Toxoplasma protein are sick and dying , and that their altered electrical activity reflects this unhealthy state . Understanding how Toxoplasma parasites persist in the brain has the potential to reveal new targets for treating symptomatic infections . It could even provide new possibilities for targeting the inflammation that drives many other neurological diseases . Harnessing this potential will require finding out why Toxoplasma gondii infects specific brain regions and why most neurons that directly interact with the parasite die .