A three-dimensional immunofluorescence atlas of the brain of the hackled-orb weaver spider, Uloborus diversus

Spiders display fascinating behaviors, perhaps none more famous than web weaving. Yet little is known about the neural processes underlying this skill. A necessary step toward unravelling this enigma is the development of a three-dimensional brain atlas of a web-building spider.

To date, spider brain anatomy has been studied primarily in species that do not build webs. Moreover, most previous work involved thin tissue sections stained to highlight specific structures. These sliced views can often be discontinuous, making it difficult to relate information across different stains and brain regions.

To overcome these limitations, Artiushin, Corver and Gordus employed an antibody-based staining method that preserves intact brains for confocal microscopy. This approach enabled the creation of a detailed, three-dimensional anatomical map. The researchers further used a computational approach to align multiple scans into a single reference volume, allowing comparative overlays of different neuron classes. This strategy made it possible to identify areas known from other spiders, as well as new features that had not been previously documented.

Using this atlas, Artiushin et al. described new details of established brain regions – including neural circuits containing major neurotransmitters and neuromodulators. They also identified previously unknown brain structures, such as a region they termed the tonsillar neuropil, as well as a potential protocerebral bridge.

This work represents the first standardized brain atlas for any spider species. While earlier studies have imaged individual neuronal populations across disparate species and brain regions, this atlas provides the most comprehensive view to date within a single species. The newly identified brain regions offer promising candidates for future functional studies and may play roles in the intricate process of web building. More broadly, these findings advance our understanding of spider neuroanatomy and will be of interest to comparative and evolutionary studies of brain structure across animals.

A brain atlas provides a foundation for functional studies that link neuronal activity to behavior. Web building in spiders is a complex, innate form of animal construction that unfolds over multiple hours in distinct stages, making it a compelling model system for investigating the neural basis of extended, sequential behaviors.

Brain atlases are essential tools for neuroscience in model organisms – ranging from neuropil annotations (Jundi and Heinze, 2020), to neuronal subtype and transcriptional expression pattern atlases (Zhang et al., 2023), to ultrastructural connectivity maps (Cook et al., 2019; Chua et al., 2023; Dorkenwald et al., 2024; Yim et al., 2024; Cook et al., 2025; Verasztó et al., 2025; White et al., 1986; Winding et al., 2023). In recent years, three-dimensional atlases of major neuropil structures have also been created for non-canonical arthropod study species (Brenneis, 2022), including a number of insects (Adden et al., 2020; Althaus et al., 2022; Dreyer et al., 2010) and spiders (Steinhoff et al., 2017; Steinhoff et al., 2024).

The hackled-orb weaver spider, Uloborus diversus (Eberhard, 1972), is an emerging model system for the study of orb-web building in spiders (Corver et al., 2021; Miller et al., 2022), whose central nervous system has yet to be investigated. To date, the majority of studies of the spider central nervous system have been performed in one de facto model species, Cupiennius salei, a cursorial spider which hunts without building webs for prey capture (Babu and Barth, 1984). While isolated anatomical treatments exist for orb weavers and other web-based spiders (Long, 2021; Hwang et al., 2015; Long, 2016; Moon and Tillinghast, 2013; Park et al., 2013; Rivera‐Quiroz and Miller, 2022; Becherer and Schmid, 1999; Steinhoff et al., 2024; Wegerhoff and Breidbach, 1995; Weltzien and Barth, 1991), the preponderance of C. salei literature is even starker when considering examinations beyond general neuronal stains, where C. salei is essentially the only spider species in which the expression pattern of more than a single neurotransmitter has been broadly mapped (Becherer and Schmid, 1999; Fabian-Fine et al., 1999; Fabian-Fine et al., 2015; Fabian-Fine et al., 2017; Loesel et al., 2011; Schmid and Becherer, 1996; Schmid and Duncker, 1993; Senior et al., 2020; Seyfarth et al., 1990; Seyfarth et al., 1993; Tarr et al., 2019). Furthermore, the current understanding of spider brain anatomy is substantially based on tissue slice analysis, which can provide exceptional detail and avoid damaging superficial brain structures, but has the disadvantage of being often limited in completeness by the planes which authors chose to exhibit.

Given that the substantial behavioral adaptation of web-building may be reflected in the presence of necessary brain structures and distinct underlying neuronal circuitry, an important step in understanding the basis of this behavior is to have a detailed, foundational architecture of a nervous system which generates it. We created a three-dimensional immunofluorescence atlas of major neurotransmitter and neuromodulator populations for U. diversus, using whole-mounted synganglia. Using immunostaining against the presynaptic marker, synapsin, we assembled a standard, full volume of the U. diversus synganglion onto which specific neurosignaling molecule expression patterns were aligned. These include markers for classical neurotransmitters (GABA and acetylcholine), neuromodulators (dopamine, serotonin, and octopamine/tyramine), and several neuropeptides (allatostatin A, proctolin, CCAP, and FMRFamide). These volumes provide comprehensive and comparable detail throughout the synganglion, in both undifferentiated and established regions – such as the arcuate body, whose layers become distinguishable through the use of neurosignaling molecule co-stains. We further identify several previously undescribed neuropils in the supraesophageal ganglion, and the neuronal subtype populations whose specific expression demarcates them.

The central nervous system of spiders is distinctive among arthropods for its compressed nature. Residing within the prosoma, the central nervous system, or synganglion, as it has been called (Steinhoff et al., 2017), is comprised of two major divisions named in reference to the esophageal passage traveling between them – the subesophageal mass, comprised primarily of motor and sensory interneurons and comparable to the ventral nerve cord in insects, and the supraesophageal mass, containing the higher-order integration centers (Figure 1A). The divisions of the synganglion have also been described in regard to the dorsal-most protocerebrum (containing the optic, arcuate, and mushroom body [MB] neuropils), a deutocerebrum (comprised of the cheliceral neuropil [ChN] and esophagus), and the dorsal and posterior fused postoral ganglia (including the leg, pedipalp, and opisthosomal neuropils [OpN]) (Long, 2021; Steinhoff et al., 2024). Consistent with general arthropod nervous system morphology, the neuronal somata are found superficially (Figure 1B), while the internal structure of the brain is comprised primarily of neuropil.

Figure 1 with 3 supplements see all

Download asset Open asset

To create a standard, three-dimensional brain atlas framework for U. diversus, we used elastix (Klein et al., 2010; Shamonin, 2014) to register and align multiple confocal image volumes of whole-mounted brain immunostains for the presynaptic marker, synapsin (3C11 antibody, DSHB). By averaging aligned image volumes (n = 6), we generated a consistent reference volume to use for further registration of more defined antibody targets, while revealing the patterns of neuropil structure of the U. diversus central nervous system, contiguously throughout the sub- and supraesophageal regions (Figure 1—video 1: synapsin), which we describe beginning at the ventral end and traveling dorsally.

Using anti-synapsin immunoreactivity as a common reference channel for all immunostains enabled application of the derived transformation matrix to respective co-stains for each brain sample, allowing incorporation of these channels into the standard brain, amounting to a total of eight neurosignaling population immunostains in addition to anti-synapsin and DAPI (nuclear marker) unified in the current atlas. In this way, even though non-synapsin antibody targets were applied to separate brains, alignment of the synapsin channel to the reference enabled comparative analysis of co-expression of several antibody targets within the same reference volume.

We have collected standard brain-aligned confocal volumes for specific neurotransmitter and neuromodulator expressing populations, including cholinergic (anti-ChAT), serotonergic (anti-5-HT), and octopaminergic/tyraminergic (anti-TDC2) (Figure 1—video 2: neurotransmitters and neuromodulators). Neuropeptidergic expression patterns have been revealed for proctolin, allatostatin A, CCAP, and FMRFamide (Figure 1—video 3: neuropeptides) – whose functional significance remains unknown, but which have been previously imaged in spiders (Loesel et al., 2011; Breidbach et al., 1995). These aligned volumes have been made available via an online repository (see Data availability).

GABAergic expression pattern (anti-GAD) is aligned and reported, but of a more limited utility as the antibody signal penetration is limited to the periphery of the tissue. Certain target populations, such as dopaminergic neuron expression patterns (anti-tyrosine hydroxylase), are also provided, although we were not able to align the volume to the standard brain due to incompatibility of optimal staining conditions between the required antibodies. The use of other antisera, such as for β-Tubulin3 and horseradish peroxidase (HRP), is also demonstrated, but without alignment to the standard brain.

Subesophageal (fused postoral ganglia) neuropils

Leg neuropils

The leg neuropils (LNs) are the first and most apparent neuropil structures to appear in the ventral subesophageal mass and are present throughout most of the horizontal planes of this mass (until ~z400, standard brain) (Figure 2A; Figure 2—video 1; Figure 2—video 2; Figure 2—video 3). Their structure and innervation is generally consistent for all legs, with the only discernible difference being that LN1 are larger than those of the other legs, allowing for innervation patterns to be more easily characterized.

Figure 2 with 3 supplements see all

Download asset Open asset

Most notably in the neuromeres of Legs I (anterior), the serotonergic (anti-5-HT) innervation in the limb neuroarchitecture appears to be supplied in two roughly equal halves, filling the periphery and leaving an area dark of immunoreactivity within (Figure 2Bi – dotted boundary). The anterior half of the innervation appears to be supplied from the medial branch of the ‘dorsal-most tract’ (as referenced by Auletta et al., 2020). Dopaminergic signal (anti-TH) is more expansive, evenly filling each LN with a mesh-like network of TH+ varicosities (Figure 2D). Unlike the approximately uniform innervation produced by dopaminergic neurons, the pattern in TDC2 staining is notably different. The anterior side of each neuromere contains a patch of continuous, diffuse, and more lightly stained immunoreactivity, while on each posterior side, there is a swath of brightly reactive, sparse puncta (Figure 2Bii). Cholinergic ChAT immunoreactivity is punctate and broadly filling of the neuropil at the ventral end (Figure 2Biii) and more limited to the medial portion on the dorsal end (Figure 2Ciii).

AstA immunoreactivity has a distinctive pattern within the leg neuromeres, showing robust varicosities but only in the dorsal–posterior portion of neuropil (Figure 2Cvii). This innervation appears to be supplied from the lateral branches of the centro-lateral tract. A similar pattern regarding expression only in the posterior aspect of the neuropil is observed for proctolin immunoreactivity (Figure 2Civ). CCAP immunoreactivity within the LNs is predominantly in the posterior halves, where sparse puncta are evenly distributed (Figure 2B, Cvi). FMRFamide immunoreactivity is also evident in the LNs (Figure 2B, Cv).

Opisthosomal neuropil

The OpN is found at the posterior and dorsal-most extension of the subesophageal mass (Figure 3A; Figure 3—videos 1–3). The region of the OpN has also been called the abdominal ganglia (Foelix, 2025), and tapers posteriorly to give way to the cauda equina, a nerve bundle composed of multiple paired tracts which innervate organs within the opisthosoma, such as the gonads, book lungs, and spinnerets (Gonzalez-Fernandez and Sherman, 1984). Within the OpN, there are a number of antero-posterior longitudinal tracts, while a ladder-like appearance of medio-laterally running tracts in the ⍺-synapsin channel can also be observed (Figure 3B, C). Posteriorly traveling tracts also diverge laterally to follow the circumference of the OpN (Figure 3B–D).

Figure 3 with 3 supplements see all

Download asset Open asset

Particular OpN features are revealed by a number of specific immunostains, including the monoamines octopamine/tyramine (TDC2 immunoreactivity), as well as neuropeptides such as proctolin and allatostatin A. TDC2 immunoreactivity displays an intricate pattern within the OpN. At the ventral anterior end, two triangular formations of puncta (Figure 3Bii – brace) abut the input of a string of varicosities on each lateral side, which then become heavier and continue to outline the boundary of the OpN (Figure 3Bii – arrow). The lateral perimeter tracts are likewise revealed by ChAT and proctolin immunoreactivity (Figure 3B, Ciii, iv) as well as by fibers from dopaminergic populations (anti-TH) (Figure 3D).

Within the interior, fibers resembling spokes emanate to a ring-like midline where there appears to be a small decussation and a thicker bridge structure joining lateral segments which travel in the anterior–posterior direction. The fine neurites projecting to the center of the OpN as seen for TDC2 immunoreactivity are also apparent for proctolin immunoreactivity (Figure 3Bii, iv – arrowhead). Proctolin signal also reveals a more posterior and dorsal crossing-over point (Figure 3Civ – arrow). Intense boutons line a tract running parallel to the midline, while also giving rise dorsally and laterally to a ladder-like structure of projections in the anterior–posterior direction, which can best be seen with octopaminergic/tyraminergic innervation (TDC2+) (Figure 3Cii – brace) and dopaminergic signal (TH+) (Figure 3D).

AstA+ immunoreactivity is also found in the OpN, with a more confined medial density in the dorso-posterior section, a pattern generally shared with 5HT and proctolin immunoreactivity (Figure 3Cvii). Ample FMRFamide signal is seen within the opisthosomal neuromere, but its signal is more uniform than the other targets investigated (Figure 3B, Cv).

Pedipalpal neuropil

As the LNs begin to diminish in the dorsal region of the subesophageal mass, a smaller antero-lateral synaptic density becomes apparent, representing the pedipalpal neuropil (PdN) (Figure 4A; Figure 4—videos 1 and 2), which serves the anterior-facing pedipalp appendages. Immunoreactivity for ChAT (acetylcholine) and TDC2 (octopamine/tyramine) is strongest among the antisera tested for the PdN, showing punctate expression which does not extend to fill the anterior portion of the neuropil (Figure 4Biii and ii, respectively). No additional structural features were evident within the PdN. Little to no appreciable immunoreactivity was seen for allatostatin A or CCAP antisera in our defined bounds of the PdN. This was also true of proctolin immunoreactivity, although signal is present in an immediately medial, adjacent region (Figure 4Biv), which is also highlighted by 5-HT immunoreactivity (see z275 in atlas). This area appears to be supplied by at least two anteriorly located proctolin+ somata (Figure 4Biv – arrow).

Figure 4 with 6 supplements see all

Download asset Open asset

Supraesophageal (Protocerebral) neuropils

Ventral features

The supraesophageal mass begins dorsally to the closure of the esophageal passage, and in its ventral-most planes, can be subdivided anteriorly and posteriorly into two sectors of slightly differing anti-synapsin immunoreactivity (Figure 5A–C – dashed perimeter, anti-synapsin). We will refer to these regions as the anterior and posterior stalk, in order to enable discussion of smaller features of various specific neurostains found within their perimeter. Lacking any further knowledge, these names are not currently intended to suggest a cohesive form or function for the structures within their bounds.

Figure 5 with 2 supplements see all

Download asset Open asset

The esophageal passage is bridged at the anterior side by a region named the stomodeal bridge (StB) (Steinhoff et al., 2017; Figure 5A – brace). A bridge structure also exists at the posterior end, where additional undifferentiated synaptic density is flanking (Figure 5B – brace with asterisk). Within these planes, a protocerebral tract (PCT) is essentially parallel to the ventro-dorsal axis and appears as twin, dense nodes rising in the central burgeoning protocerebrum (Figure 5B, C).

5-HT immunoreactivity is prominent in the posterior bridging area dorsal to the esophageal passage (Figure 5B, Ci, Figure 5—video 1), as well as the laterally adjacent tissue, and not as apparent in the anterior StB. 5-HT immunoreactivity is otherwise weak within the stalk regions. TDC2 immunoreactivity is prominent in the StB and adjacent areas, and at this plane, two lateral bands of immunoreactivity appear which do not correspond to a clear demarcation in the synapsin channel (Figure 5Biii – arrow, Figure 5—video 1).

Dorsally, octopaminergic/tyraminergic (TDC2+) signal is prominent along an antero-lateral stretch marking the boundary of what we define as the posterior stalk, a perimeter also visible in the synapsin channel (Figure 5Ciii). Concentrations of TDC2+ immunoreactivity are also apparent centrally, both anterior and lateral to the PCTs (Figure 5Ciii – arrows, Figure 5—video 1).

Similar to what has been described for M. muscosa as the StB (Steinhoff et al., 2017), the area adjacent to the esophagus on the anterior side has immunoreactivity to allatostatin A, although the actual bridge which crosses the midline is modest, with thin representation in the posterior commissure (Figure 5Av – arrowhead, Figure 5—video 2).

In U. diversus, strong AstA immunoreactivity is present on the posterior side of where the esophagus closes, in the posterior stalk area (Figure 5Cv). The posterior region adjacent to the midline, previously highlighted with 5HT immunoreactivity, also shows partial AstA+ innervation, displaying a unique oxbow type pattern (Figure 5Cv – arrow, Figure 5—video 2). There is also a patch of AstA+ immunoreactivity in the central area of the anterior stalk (Figure 5Cv – brace).

On the posterior edge of the StB, there is a thin Proc+ commissure, while the anterior edge of the StB is highlighted by a bolder vein of varicosities (Figure 5Aiv – arrows). Proctolin signal is present adjacent to the midline around the posterior bridging area, shared with 5-HT, AstA, and TDC2+ immunoreactivity, and together with the commissure evident by synapsin staining, forms a circular pattern (Figure 5Biv – brace). This circular pattern of immunoreactivity is also visible by 5-HT+ immunoreactivity (Figure 5Bi), as well as TDC2+ innervation, though more clearly seen further dorsally (Figure 5Ciii) for this channel. There is also proctolin immunoreactivity seen centrally, medial to where the PCT is ascending (Figure 5B, Civ).

ChAT+ immunoreactivity is diffusely present throughout the stalk regions (Figure 5Cii), and unlike the other antisera, co-stains the posterior aspect of the PCT, which is prominently visible with synapsin immunoreactivity (Figure 5B, Cii – arrowheads and dashed circles).

Hagstone neuropil and mid-supraesophageal features

We will define the hagstone neuropil as a central, midline adjacent, paired structure, whose ventral bounds share the same plane as the appearance of a prominent protocerebral commissure (standard brain, ~z = 540), and which continues dorsally until the formation of the MB bridge (standard brain, ~z = 615). This approximately kidney-bean-shaped structure is pierced by a circular spot lacking synapsin immunoreactivity (Figure 6A, C, D – dashed outline, Figure 6—videos 1–3) – which may be indicative of a space occupied by a fiber tract, tracheal passageway, and/or potentially glia.

Figure 6 with 3 supplements see all

Download asset Open asset

The hagstone neuropil, as well as other prominent features of the mid-central protocerebrum, are most clearly defined by serotonergic immunoreactivity. Serotonergic fibers form a distinctive circular tract pattern around the midline of the supraesophageal ganglion (Figure 6Bi – brace), not as clearly seen with any other neuronal-subtype stain. The semi-circular tracts bend medially (Figure 6Bi), before a fusion of seemingly all three directions is seen immediately dorsal (Figure 6C). The hagstone neuropil is essentially completely filled with serotonergic immunoreactivity, matching the outline defined in the anti-synapsin channel (Figure 6C, Di).

On the posterior end is a diffuse, arching band of varicosities, with fiber tracts at the midline, resembling an umbrella-like form (Figure 6C, Di – brace, anti-5-HT). While the umbrella-like band formation is not distinct enough to be reliably annotated from synapsin immunoreactivity, we find that signal within this formation is also visible in other neuromodulator and neuropeptide immunostains.

TDC2 immunoreactivity is also present in the umbrella-like posterior region innervated by 5-HT (Figure 6Ciii – brace, Figure 6—video 2), and sparser puncta within the bounds of the hagstone neuropil. Proctolin immunoreactivity is present in the posterior, midline-spanning umbrella structure observed for 5-HT and TDC2, as well as fine varicosities in the hagstone neuropil (Figure 6Dii). This is likewise the case for AstA immunoreactivity in the hagstone neuropil, while posteriorly, there is abundant AstA+ signal which partially is coincident with the umbrella-like band. Posterior to the cup-shaped synaptic density formed by the MB hafts continuing with the rest of the MB, is a crescent of proctolin immunoreactivity (Figure 6Cii – brace), which also appears to be present in C. salei (Becherer and Schmid, 1999). ChAT immunoreactivity, as before, is found broadly, including within the hagstone neuropil, though no greater structure is discernible from this channel in the mid-supra otherwise. It is difficult to ascertain to what degree the innervation in this region is continuous with that of dorsally located features.

Mushroom bodies

The mushroom bodies (MBs) of U. diversus (Figure 7A, B; Figure 7—video 1) tend to show the most robust synapsin immunoreactivity of all structures in the supraesophageal mass (Figure 7B, maximum intensity projection), indicating a great degree of synaptic density. While web-building species have been reported to have simplified (Steinhoff et al., 2024) or even entirely absent mushroom bodies (Hanström, 1928; Long, 2016; Long, 2021; Steinhoff et al., 2024) although notable exceptions have also been observed such as D. spinosa and A. trifasciata (Long, 2016), these structures are present in U. diversus and retain the complete form seen in more visually reliant species (Steinhoff et al., 2017; Steinhoff et al., 2024), even if they are smaller relative to the protocerebrum as a whole (Figure 7A–C).

Figure 7 with 1 supplement see all

Download asset Open asset

U. diversus MBs display a haft, body, and head region, with the two pairs connected by a bridge (Figure 7A–C). Synapsin immunoreactivity is modest within the bridge region, whose true thickness is better visualized with staining for βTubulin3 (Figure 7D). Despite the strong synapsin immunoreactivity in the MBs, we surprisingly did not see co-expression with most of our specific neurosignaling molecule antibodies. This pattern is also reflected in the extant spider literature, with a single study showing immunoreactivity in the mushroom bodies of C. salei for anti-GAD and anti-proctolin staining (Becherer and Schmid, 1999). A particularly intensely stained MB bridge was also seen in A. sclopetarius using a non-antisera stain of acetylcholinesterase activity (Meyer and Pospiech, 1977). In our hands, only anti-allatostatin A staining showed co-expression throughout the MB (Figure 7E). Although difficult to trace the source, it appears the hafts are innervated from the posterior side (Figure 7F – arrows). By βTubulin3 immunoreactivity, we observe two tracts which straddle the MB hafts as they descend from the dorsal somata layer (Figure 7G). Finer neurites are not distinguishable in the βTub3 immunoreactivity, but it is plausible that the AstA+ neurites entering the MB hafts might stem from the medial of these two tracts.

Babu and Barth, 1984 described the protocerebro-dorsal tract as providing input to the hafts of the mushroom bodies. The connection of this tract to the MB hafts is not apparent by synapsin or βTubulin3 immunoreactivity in U. diversus, which was likewise the case with silver staining for P. amentata, M. muscosa, A. bruennichi, and P. tepidariorum (Steinhoff et al., 2024).

The antero-dorsal input to the MB heads, representing the secondary eye pathway (Strausfeld and Barth, 1993), is much more conspicuous and has received considerable treatment within the literature. The MB heads are sometimes referred to as the third-order visual neuropil in this pathway, with the ample parallel fibers which give this structure its shape arising from globuli cells which cap the MB head.

The globuli cells are not distinguishable from the surrounding nuclei by DAPI signal, but can potentially be discerned through specific neurosignaling molecule immunostains. We find the cluster of cells closely associated with the MB heads is revealed by ChAT immunoreactivity, and to a lesser extent by GAD immunoreactivity, suggesting they represent cholinergic and GABAergic populations, respectively (Figure 7H, I – arrow(s)). Globuli cells in C. salei have previously been shown to be ChAT+ (Fabian-Fine et al., 2017). By βTubulin3 staining, we also observed a trident of tracts feeding into the dorsal aspect of the MB head (Figure 7G).

Visual system

U. diversus, like many orb weavers, builds its web at night and can do so in essentially complete darkness in laboratory conditions, suggesting that vision is expendable to much of the spider’s behavioral repertoire (Eberhard, 1971). Web-building spiders are considered to have poorer vision than spiders which depend on sight to capture prey, which is reflected in their diminished optic neuropils and tract pathways (Long, 2021; Rivera‐Quiroz and Miller, 2022; Steinhoff et al., 2024).

The number and particularly size and spatial arrangement of spider eyes is variable and characteristic across species, but most commonly, spiders have eight eyes which give rise to two visual system pathways, the principal and secondary eye pathways (Strausfeld and Barth, 1993; Strausfeld and Barth, 1993; Long, 2021; Steinhoff et al., 2024). The pair of anterior medial eyes is also known as the principal eyes which innervate the principal pathway terminating primarily in the arcuate body. The remaining pairs are the six secondary eyes, including the anterior lateral eyes, posterior medial eyes, and posterior lateral eyes, whose projections form the secondary eye pathway. U. diversus has eight eyes which are approximately equal in size.

Relative to cursorial species (Babu and Barth, 1984; Steinhoff et al., 2017; Steinhoff et al., 2024), in U. diversus the anterior extensions of the protocerebrum containing the first and second-order optic neuropils are considerably thinner and not as extensively fused with the continuous neuropil of the supraesophageal mass, thus being prone to separating during dissection. Consequently, neither the primary nor secondary visual pathway neuropils appear reliably enough in the anti-synapsin volumes to be apparent in the averaged standard brain representation, but nevertheless, these structures are exhibited in various individual preparations. The optic neuropils in U. diversus tended to show weaker synapsin immunoreactivity but were clearly seen with antisera to HRP (Figure 8A).

Figure 8

Download asset Open asset

As in other species, the secondary pathway is larger (Figure 8B), lifting away anteriodorsally from the zone of the MB heads. This continuity can be seen from the sliced three-dimensional maximum intensity projection of synapsin (Figure 8B – brace). The primary pathway is diminutive in U. diversus and emerges as a bulbous shape at the dorsal-most end of the brain through a field of somata (Figure 8A).

Previous reports have used GABA (Becherer and Schmid, 1999), histamine (Schmid and Duncker, 1993), dopamine (Auletta et al., 2020), CCAP (Loesel et al., 2011), and FMRFamide (Becherer and Schmid, 1999) to reveal the successive neuropils of the visual pathways. As noted above, the only features within the optic pathway for which we observed neurosignaling molecule immunoreactivity were the globuli cells with GAD and ChAT staining. It is possible that targets for which we could not acquire an effective antisera, such as histamine, could be revelatory of the optic lamellae and other visual pathway structures, as they have been for C. salei (Schmid and Becherer, 1999; Schmid and Duncker, 1993). Specific compartments of the pathways, such as the medulla or lamellae, could not be confidently discerned with any preparation.

Tonsillar neuropil

Within the historically non-descript central protocerebrum, we observed a synaptically dense neuropil structure in U. diversus. Beginning in the planes dorsal to the mushroom bodies, this paired structure is positioned directly on either side of the midline and is centrally located. Each half has an approximately ovoid appearance, particularly at the anterio-dorsal end, while being bridged at the posterior aspect. Between the two halves, at the midline, is a furrow which is negative for synapsin immunoreactivity, giving this neuropil, in conjunction with the synapsin-negative zone, a likeness to tonsils when viewed from the horizontal optical planes (Figure 9A, C– α-synapsin, Figure 9—videos 1–3) – hence our referral to this structure as the tonsillar neuropil.

Figure 9 with 3 supplements see all

Download asset Open asset

In individual anti-synapsin stains, a fiber tract traveling laterally adjoins this neuropil in the more dorsal–posterior portions. By tubulin immunoreactivity, it appears to bifurcate the structure below the bridge in the dorsal portion (Figure 9B). A subset of antisera for specific neuronal populations is instrumental in confirming this neuropil, as their immunoreactivity is circumscribed by its boundaries, with little neighboring signal to obscure the distinction (Figure 9C). Most representative among these is serotonergic immunoreactivity, exhibiting fine varicosities which neatly fill the area (Figure 9Ci, vi). TDC2+ signal is also prominent in this neuropil Figure 9Cii, vii – the relatively heavier terminals appear stronger on the periphery, and when viewed in alignment with the 5-HT channel, resemble a division of compartments, most notably in the ovoid regions where serotonin is found in an internal, core pattern, with octopaminergic/tyraminergic signal as a shell.

There may also be a division in the anterior–posterior dimension as allatostatin A immunoreactivity is more pronounced in the posterior bridging region and sparsely punctuated in the anterior ovoid zones (Figure 9Civ, ix – α-AstA), while proctolin immunoreactivity is limited to the posterior region (Figure 9Ciii, viii – α-proctolin). FMRFamide immunostaining is diffusely present, particularly in the anterior–dorsal portions of the tonsillar neuropil (Figure 9Cv, x), but this is amidst broadly saturated signal from this antibody throughout the protocerebrum.

Protocerebral bridge

Originating antero-laterally and progressing posterior-medially through the ascending dorsal planes of the supraesophageal ganglion is a banded neuropil structure which we will designate as the protocerebral bridge (PCB). Wider in the lateral aspect, the structure tapers toward the medial end with the thinnest, midline-crossing component only being apparent in specific neuronal subpopulation stains. This is the dorsal-most neuropil seen in the interior of the protocerebrum before reaching the dense cap of somata (Figure 10; Figure 10—videos 1–3).

Figure 10 with 3 supplements see all

Download asset Open asset

As with the previous neuropils, only a subset of antisera shows immunoreactivity within this structure. The most filling is GABAergic immunoreactivity (by anti-GAD stain) which defines a nearly complete swath of the neuropil with dense signal (Figure 10Bi, iv – α-GAD). Comparing further neuronal subtype preparations reveals that the PCB has a layered structure. TDC2 immunoreactivity is pronounced throughout the length of the protocerebral bridge, displaying heavy chains of puncta on the posterior edge of the bridge (Figure 10Bii, vii – α-TDC2). Proctolin immunoreactivity forms a tight, thinner band, primarily at the medial end of the neuropil, comprised of fine puncta and is centrally located among the layers (Figure 10Biii, viii – α-proctolin). The cholinergic pattern is a distinct thin layer on the anterior and posterior edge, most clearly visible on the lateral portion of the PCB (Figure 10Biv, ix – α-ChAT). A faint section of AstA immunoreactivity was also seen (Figure 10Bv, x – α-AstA).

Posterior to the PCB is an arching string of varicosities which reaches its apex just before the appearance of the dorsal arcuate body layer. We refer to this as the dorsal protocerebral commissure (Figure 10B – arrows) and the neuronal subtype populations which show PCB expression tend to also innervate this commissure. The strongest of these are octopaminergic/tyraminergic neurons (anti-TDC2) and proctolin+ neurons (anti-proctolin) (Figure 10B – α-TDC2, α-proctolin). This pathway separates from the posterior–lateral contour of the PCB, where there is an approximately triangular expansion of immunoreactivity before traveling medially and passing just posterior to the edge of the tonsillar neuropil. A similar pattern is also seen for allatostatin A (Figure 10B – α-AstA). Cholinergic immunoreactivity is also apparent in the posterior arch, but is more subtle and in a single layer in the anterior domain, less comparable to that of TDC2, proctolin, and allatostatin A.

Arcuate body

The arcuate body (AB) is a prominent neuropil structure found in all spider species whose central nervous system anatomy has been examined closely to date. Residing in the dorso-posterior aspect of the supraesophageal mass, this solitary crescent-shaped structure has been recognized as having at least two broad divisions, the ventral and dorsal lobes (Hanström, 1928; Loesel et al., 2011; Steinhoff et al., 2017).

In U. diversus, at the grossest level, we likewise observe two lobes of the arcuate body, which we will refer to as the ventral (ABv) and dorsal (ABd) lobe (Figure 11A, B; Figure 11—videos 1–3). Though largely coincident in the dorso-ventral axis, the ventral arcuate body somewhat envelopes the dorsal arcuate lobe, hence appearing first from the ventral direction and lingering posteriorly on the dorsal side, with only a smaller part of the dorsal arcuate body protruding independently beyond the ABv at the dorsal end (Figure 11B; Figure 11—video 1).

Figure 11 with 3 supplements see all

Download asset Open asset

Each lobe (ABv and ABd) can be further subdivided into two sub-lobes or layers – a posterior (posABv and posABd) and anterior (antABv and antABd) section, making a total of four units (Figure 11C). The sublayers of the arcuate body lobes are distinguishable by immunostaining for specific neuronal subpopulations that differentially innervate the layers (Figures 11D and 12). By examining these expression patterns, another tier of complexity can be appreciated, as each of these sublayers (posABv, antAbv, posABd, and antABd) can be further subdivided into two or even three aspects, depending on the antisera used.

Figure 12

Download asset Open asset

Posterior to the arcuate body is a crest of somata which has been previously referred to as the posterior cell layer (PCL) (Fabian-Fine et al., 2017). Neurons of the PCL send their projections anteriorly through the ventral arcuate layers, as revealed by immunostaining for βTubulin3 in conjunction with synapsin (Figure 11E). The fibers successively appear more medially as one progresses further dorsally in the arcuate lobes, with certain tracts being thicker than others. Hill noted the presence of tracts running through the arcuate body to join the PCDt in the jumping spider, P. johnsoni (Hill, 1975).

The innervation pattern of a given neuronal subpopulation in a layer of the arcuate body is not a general delineation of the structure of that layer, as different transmitter populations can display distinct expression patterns within the same layer. An example is the dorsal arcuate body (ABd), where TDC2 immunoreactivity shows a prominent columnar, flagstone innervation, while proctolin has a sparse field of fine puncta in the same layer (Figure 11F). Even though the arcuate body is organized in lateral bands, columnar organization can also be observed in several layers. For example, a series of discernible units of allatostatin A+ immunoreactivity are seen in the posABv layer, suggesting the columnar organization which is present, but generally obscured by the density of staining (Figure 11G).

There is a diversity of layering patterns (Figure 12), but some basic motifs emerge. Innervation can be partial, as in filling a single sublayer (anterior or posterior) of a lobe, or complete throughout the lobe, taking on a saturated appearance, a meshwork of neurites, or a sparse field of puncta. The space between marked sublayers may have finer neurite connections which have been described as palisade-like (Loesel et al., 2011). At the dorsal end of the ventral arcuate body (ABv), heavy garland-like varicosities may form, in certain examples (α-proctolin, α-ChAT, Figure 12 – arrowheads, on respective subpanels) appearing as disjointed units, suggestive of an undergirding column. More prevalent in the dorsal arcuate, a robust networking of thicker immunoreactive fibers weaves between roughly trapezoidal signal-negative areas (α-5-HT, α-TDC2, Figure 12 – braces, on respective subpanels), resembling a flagstone pathway. Detailed descriptions of arcuate body layer projection patterns (Figure 12) and comparisons to other spider species are found below in the respective subsections for each neurosignaling molecule.

GABA

In the ventral lobe of the arcuate body, GAD immunoreactivity is faint and difficult to discern from bleed-through or background; however, we see a clear illustration of how neurites stemming from the somata of the PCL extend through the arcuate body layers (Figure 12i – α-GAD, ventral – arrows). At the edge of the posterior ABd is a layer of diffuse immunoreactivity (Figure 12ii – α-GAD, dorsal – arrow). Moving anteriorly, this is followed by a thin layer of puncta (Figure 12ii – α-GAD, dorsal – arrowhead), which may be connected through minute projections to the next layer which is thicker than the first (Figure 12ii – α-GAD, dorsal – brace). The pattern and relative proportions of the layers are broadly similar in C. salei (Becherer and Schmid, 1999; Fabian-Fine et al., 2015), where the anterior-most layer in the dorsal arcuate body is also the thickest, albeit less densely stained in our example.

Acetylcholine

In the arcuate body, cholinergic signal is predominantly found in the ventral arcuate body lobe (Figure 12iii – α-ChAT, ventral). Within the ventral side of this lobe, cholinergic immunoreactivity forms fine puncta which completely fill the anterior sub-layer of this lobe (Figure 12iii – α-ChAT, ventral – brace). Toward the dorsal end of this lobe, the punctate immunoreactivity forms heavier beaded varicosities (Figure 12iv – α-ChAT, dorsal – arrowhead). Midway, there are faint column-like expression patterns joining from a thin layer within the posterior ventral AB (pABv). A broader layer is innervated on the anterior side of the dorsal lobe (Figure 12iv – α-ChAT, dorsal – brace), a sparser example of the flagstone-type patterning seen more clearly with α-TDC2 immunoreactivity, below. Acetylcholinesterase activity has been visualized through acetylthiocholine iodide staining in cursorial and web-building species (Meyer and Idel, 1977; Meyer and Pospiech, 1977). The lycosid T. spinipalpis and salticid M. muscosa appear to show at least two broad layers of cholinergic signal, similar to U. diversus (Meyer and Idel, 1977). The orb-web-building araneid A. sclopetarius had relatively diminished arcuate body signal compared to the central protocerebrum of the cursorial species with this preparation (Meyer and Pospiech, 1977), and the multilayered signal was not as clear. Since it can be difficult to judge which arcuate body lobes are on display from a single slice, it is also possible that the diminished signal in the anterior layer for A. sclopetarius might be consistent with U. diversus patterning, if that is indeed the dorsal arcuate body lobe.

Dopamine

Within the arcuate body, TH immunoreactivity occupies a single layer in the posterior aspect of the dorsal lobe (Figure 12vi – α-TH, dorsal – arrow), supplied by thin and sparse neurites stretching from the anteriorly located tracts. This single layer of punctate terminals with anteriorly branching projections is consistent with both H. lenta and P. regius (Auletta et al., 2020), but otherwise, the U. diversus dopaminergic arcuate body layering appears simpler and more comparable to the jumping spider, P. regius, due to lacking the additional wispy immunoreactivity in anterior layers as in H. lenta.

For the wolf spider, H. lenta, TH labeling reveals densely stained first and second-order optic neuropils (Auletta et al., 2020). In contrast, we see a stark lack of immunoreactivity in anterior regions which would be expected to contain the comparable neuropils in U. diversus.

Serotonin

Immunostaining against 5-HT in the social huntsman, Delana cancerides, shows two gross levels of immunoreactivity in the arcuate body; a wide diffuse layer of puncta, and a thinner layer bordered by dense puncta on each side, with columnar-like expression in between (Strausfeld et al., 2006). Taken together as two adjacent layers, this pattern is remarkably similar to that seen for our model species. In U. diversus, serotonergic immunoreactivity shows a faint layer in the posterior ventral arcuate lobe (Figure 12vii – α-5-HT, ventral – arrow), and an anterior ventral arcuate sublayer broadly flush with minutely fine fibers (Figure 12vii – α-5-HT, ventral – brace). The dorsal arcuate lobe displays a robust and wide immunoreactive pattern resembling the flagstone pattern, hinting at a columnar structure (Figure 12viii – α-5-HT, dorsal – brace). Innervation in this layer greatly resembles that seen for TDC2 in the same lobe (Figure 12 – α-TDC2, dorsal – brace).

Octopamine/tyramine

Both anterior and posterior sublayers of the ventral arcuate body exhibit TDC2 immunoreactivity (Figure 12ix – α-TDC2, ventral). The posterior ABv is saturated with diffuse puncta, which are also seen anteriorly, along with denser varicosities in this sublayer (Figure 12ix – α-TDC2, ventral – arrow). In the dorsal arcuate body lobe, TDC2 immunoreactivity appears prominently in the anterior sublayer (aABd), where it fully fills the span of this layer with robust staining resembling a series of flagstone-shaped columnar-like elements (Figure 12x – α-TDC2, dorsal – brace). Octopaminergic expression has been reported in the arcuate body (labeled ‘central body’ in source) (Seyfarth et al., 1993) of C. salei, where a parasagittal section shows strong immunoreactivity in the ventral portion of both arcuate body lobes. We must imagine the respective horizontal view, but it would appear by the gaps in immunoreactivity that a dorsal horizontal slice in C. salei should show three general layers of AB staining, which is essentially what we see from a dorsal plane in U. diversus.

Allatostatin A

The pattern of arcuate body innervation by AstA+ neurons is in general agreement with findings from C. salei and M. muscosa (Loesel et al., 2011; Steinhoff et al., 2017) where signal is prominent in the ventral arcuate lobe (ABv), with little staining in the dorsal arcuate (ABd) (Figure 12xi, xii – α-AstA). Concerning the sublayers of the ventral arcuate lobe, AstA immunoreactivity is seen primarily on the anterior side of the posterior ventral arcuate (posABv) and fully encompasses the anterior ventral arcuate (antABv) (Figure 12xi, xii – α-AstA – brace).

Proctolin

Proctolin immunoreactivity is evident in all lobes and layers of the arcuate body (Figure 12 – α-proctolin). In the ventral arcuate body (ABv), at the posterior end, there is a line of intense terminals (Figure 12xiii – α-proctolin, ventral – arrowhead), with more anterior layers filled with diffuse puncta. Toward the dorsal end of the ventral arcuate body (ABv), the proctolin immunoreactivity in the posterior-most layer transforms into heavy garland-like columnar varicosities extending at an anterio-dorsal angle (Figure 12xiv – α-proctolin, dorsal – arrowhead). This is in complete correspondence to the varicosities seen for this layer in C. salei (Loesel et al., 2011). In the anterior ventral arcuate (antABv), the anterior and posterior sublayers (Figure 12xiii – α-proctolin, ventral – braces) take on an intricate mesh-like form, also with smaller flagstone formations. Between these two layers are fine palisade neurites. Both sublayers of the dorsal arcuate (ABd) are also filled, but with a sparse field of fine puncta (Figure 12xiv – α-proctolin, dorsal).

CCAP

CCAP expression is strong in the posterior ventral AB layer with a fine mesh, punctate appearance which seemingly contours the columnar structures on the anterior and posterior boundaries of this layer. The anterior and posterior sublayers of the ventral AB are highlighted, with a decrease in staining within the area between the sublayers (Figure 12xv – α-CCAP, ventral – braces). In the anterior ventral AB, the anti-CCAP expression is slightly finer and more punctate than in the posterior layer. Within the dorsal AB (Figure 12xvi – α-CCAP, dorsal), only the posterior layer has appreciable expression, showing a single, finely innervated but moderately thick layer on the posterior boundary of the dorsal AB. CCAP-immunoreactive layers in U. diversus are comparable to C. salei (Loesel et al., 2011), as for both species the thickest staining layer is the most posterior one (ventral arcuate body lobe), followed anterio-dorsally by a lesser layer, and with a thinner strand of intensely immunoreactive boutons running through the more anteriorly located dorsal arcuate body lobe. CCAP immunoreactivity in two distinct layers of the arcuate body appears to be a consistent characteristic of the arcuate body lobes across arachnids, with the source also being repeatedly attributable to the ‘l’ and ‘dl’ cluster neurons (Breidbach et al., 1995). While we also see abundant CCAP+ somata, the limited expression within their projections does not allow us to confirm whether similar clusters are innervating the arcuate body layers in U. diversus.

Almost the entirety of spider central nervous system literature has been studied from tissue slices, with few examples of whole mounts (Auletta et al., 2020; Breidbach et al., 1995; Schmid and Duncker, 1993; Steinhoff et al., 2017; Steinhoff et al., 2024). Our ability to observe novel structures and make comparisons between innervation patterns was aided by whole-mount preparation and averaged brain alignment, which allowed us to compare the relative expression of targets within the same reference volume. Furthermore, imaging and alignment of many neurosignaling molecule stains in a single species were clarifying for the identification of novel structures, as a subset of stains crystallized putative boundaries. While a dozen or so species have been studied for the expression patterning of individual neurosignaling molecules (Auletta et al., 2020; Breidbach et al., 1995; Hwang et al., 2015; Moon and Tillinghast, 2013; Schmid and Becherer, 1999; Steinhoff et al., 2017; Strausfeld et al., 2006), the wandering spider, C. salei, is essentially the only species prior to the current work to have been the subject of sustained efforts to amass such neuroanatomical descriptions for most of the canonical neurotransmitters and neuromodulators. Given the utility of specific stains for understanding of neuropil structures (Figure 13, Figure 13—video 1), tracts, and other features, this atlas provides a rich source for comparative anatomy in an orb-weaving spider, U. diversus, while also extending knowledge of a number of different neurosignaling pathways for spiders at large.

Figure 13 with 1 supplement see all

Download asset Open asset

TIFF stacks of volumes presented can be viewed and downloaded from the Brain Imaging Library (BIL): https://doi.org/10.35077/ace-owl-gum. Custom code used to align volumes with elastix can be found here: https://github.com/GordusLab/Artiushin-elastix-eLife (copy archived at GordusLab, 2026).

1. 1. Artiushin G
   2. Corver A
   3. Gordus A

   (2025) Brain Image Library

   A three-dimensional immunofluorescence atlas of the brain of the hackled-orb weaver spider, Uloborus diversus.

   https://doi.org/10.35077/ace-owl-gum

1. 1. Adden A
   2. Wibrand S
   3. Pfeiffer K
   4. Warrant E
   5. Heinze S

   (2020) The brain of a nocturnal migratory insect, the Australian Bogong moth

   The Journal of Comparative Neurology 528:1942–1963.

   https://doi.org/10.1002/cne.24866

   • PubMed
   • Google Scholar
2. 1. Althaus V
   2. Jahn S
   3. Massah A
   4. Stengl M
   5. Homberg U

   (2022) 3D-atlas of the brain of the cockroach Rhyparobia maderae

   The Journal of Comparative Neurology 530:3126–3156.

   https://doi.org/10.1002/cne.25396

   • PubMed
   • Google Scholar
3. 1. Anton S
   2. Tichy H

   (1994) Hygro- and thermoreceptors in tip-pore sensilla of the tarsal organ of the spider Cupiennius salei: innervation and central projection

   Cell and Tissue Research 278:399–407.

   https://doi.org/10.1007/BF00414182

   • Google Scholar
4. 1. Aso Y
   2. Hattori D
   3. Yu Y
   4. Johnston RM
   5. Iyer NA
   6. Ngo TTB
   7. Dionne H
   8. Abbott LF
   9. Axel R
   10. Tanimoto H
   11. Rubin GM

   (2014) The neuronal architecture of the mushroom body provides a logic for associative learning

   eLife 3:e04577.

   https://doi.org/10.7554/eLife.04577

   • PubMed
   • Google Scholar
5. 1. Auletta A
   2. Rue MCP
   3. Harley CM
   4. Mesce KA

   (2020) Tyrosine hydroxylase immunolabeling reveals the distribution of catecholaminergic neurons in the central nervous systems of the spiders Hogna lenta (Araneae: Lycosidae) and Phidippus regius (Araneae: Salticidae)

   The Journal of Comparative Neurology 528:211–230.

   https://doi.org/10.1002/cne.24748

   • PubMed
   • Google Scholar
6. 1. Babu KS
   2. Barth FG

   (1984) Neuroanatomy of the central nervous system of the wandering spider, Cupiennius salei (Arachnida, Araneida)

   Zoomorphology 104:344–359.

   https://doi.org/10.1007/BF00312185

   • Google Scholar
7. 1. Becherer C
   2. Schmid A

   (1999) Distribution of γ-aminobutyric acid-, proctolin-, Periplaneta hypertrehalosaemic hormone- and FMRFamide-like immunoreactivity in the visual ganglia of the spider Cupiennius salei Keys

   Comparative Biochemistry and Physiology Part A 122:267–275.

   https://doi.org/10.1016/S1095-6433(99)00010-0

   • Google Scholar
8. 1. Breidbach O
   2. Dircksen H
   3. Wegerhoff R

   (1995) Common general morphological pattern of peptidergic neurons in the arachnid brain: crustacean cardioactive peptide-immunoreactive neurons in the protocerebrum of seven arachnid species

   Cell and Tissue Research 279:183–197.

   https://doi.org/10.1007/BF00300703

   • PubMed
   • Google Scholar
9. 1. Brenneis G

   (2022) The visual pathway in sea spiders (Pycnogonida) displays a simple serial layout with similarities to the median eye pathway in horseshoe crabs

   BMC Biology 20:27.

   https://doi.org/10.1186/s12915-021-01212-z

   • PubMed
   • Google Scholar
10. 1. Chua NJ
    2. Makarova AA
    3. Gunn P
    4. Villani S
    5. Cohen B
    6. Thasin M
    7. Wu J
    8. Shefter D
    9. Pang S
    10. Xu CS
    11. Hess HF
    12. Polilov AA
    13. Chklovskii DB

    (2023) A complete reconstruction of the early visual system of an adult insect

    Current Biology 33:4611–4623.

    https://doi.org/10.1016/j.cub.2023.09.021

    • PubMed
    • Google Scholar
11. 1. Cook SJ
    2. Jarrell TA
    3. Brittin CA
    4. Wang Y
    5. Bloniarz AE
    6. Yakovlev MA
    7. Nguyen KCQ
    8. Tang LTH
    9. Bayer EA
    10. Duerr JS
    11. Bülow HE
    12. Hobert O
    13. Hall DH
    14. Emmons SW

    (2019) Whole-animal connectomes of both Caenorhabditis elegans sexes

    Nature 571:63–71.

    https://doi.org/10.1038/s41586-019-1352-7

    • PubMed
    • Google Scholar
12. 1. Cook SJ
    2. Kalinski CA
    3. Loer CM
    4. Memar N
    5. Majeed M
    6. Stephen SR
    7. Bumbarger DJ
    8. Riebesell M
    9. Conradt B
    10. Schnabel R
    11. Sommer RJ
    12. Hobert O

    (2025) Comparative connectomics of two distantly related nematode species reveals patterns of nervous system evolution

    Science 389:eadx2143.

    https://doi.org/10.1126/science.adx2143

    • PubMed
    • Google Scholar
13. 1. Corver A
    2. Wilkerson N
    3. Miller J
    4. Gordus A

    (2021) Distinct movement patterns generate stages of spider web building

    Current Biology 31:4983–4997.

    https://doi.org/10.1016/j.cub.2021.09.030

    • PubMed
    • Google Scholar
14. 1. Doeffinger C
    2. Hartenstein V
    3. Stollewerk A

    (2010) Compartmentalization of the precheliceral neuroectoderm in the spider Cupiennius salei: development of the arcuate body, optic ganglia, and mushroom body

    The Journal of Comparative Neurology 518:2612–2632.

    https://doi.org/10.1002/cne.22355

    • PubMed
    • Google Scholar
15. 1. Dorkenwald S
    2. Matsliah A
    3. Sterling AR
    4. Schlegel P
    5. Yu SC
    6. McKellar CE
    7. Lin A
    8. Costa M
    9. Eichler K
    10. Yin Y
    11. Silversmith W
    12. Schneider-Mizell C
    13. Jordan CS
    14. Brittain D
    15. Halageri A
    16. Kuehner K
    17. Ogedengbe O
    18. Morey R
    19. Gager J
    20. Kruk K
    21. Perlman E
    22. Yang R
    23. Deutsch D
    24. Bland D
    25. Sorek M
    26. Lu R
    27. Macrina T
    28. Lee K
    29. Bae JA
    30. Mu S
    31. Nehoran B
    32. Mitchell E
    33. Popovych S
    34. Wu J
    35. Jia Z
    36. Castro MA
    37. Kemnitz N
    38. Ih D
    39. Bates AS
    40. Eckstein N
    41. Funke J
    42. Collman F
    43. Bock DD
    44. Jefferis G
    45. Seung HS
    46. Murthy M
    47. FlyWire Consortium

    (2024) Neuronal wiring diagram of an adult brain

    Nature 634:124–138.

    https://doi.org/10.1038/s41586-024-07558-y

    • PubMed
    • Google Scholar
16. 1. Dreyer D
    2. Vitt H
    3. Dippel S
    4. Goetz B
    5. El Jundi B
    6. Kollmann M
    7. Huetteroth W
    8. Schachtner J

    (2010) 3D Standard brain of the red flour beetle tribolium castaneum: A tool to study metamorphic development and adult plasticity

    Frontiers in Systems Neuroscience 4:3.

    https://doi.org/10.3389/neuro.06.003.2010

    • PubMed
    • Google Scholar
17. 1. Eberhard WG

    (1971) The ecology of the web ofUloborus diversus (Araneae: Uloboridae)

    Oecologia 6:328–342.

    https://doi.org/10.1007/BF00389107

    • PubMed
    • Google Scholar
18. 1. Eberhard WG

    (1972) The web of uloborus diversus (Araneae: Uloboridae)

    Journal of Zoology 166:4968.

    https://doi.org/10.1111/j.1469-7998.1972.tb04968.x

    • Google Scholar
19. 1. Fabian R
    2. Seyfarth EA

    (1997) Acetylcholine and histamine are transmitter candidates in identifiable mechanosensitive neurons of the spider cupiennius salei: an immunocytochemical study

    Cell and Tissue Research 287:413–423.

    https://doi.org/10.1007/s004410050766

    • PubMed
    • Google Scholar
20. 1. Fabian-Fine R
    2. Höger U
    3. Seyfarth EA
    4. Meinertzhagen IA

    (1999) Peripheral synapses at identified mechanosensory neurons in spiders: three-dimensional reconstruction and GABA immunocytochemistry

    The Journal of Neuroscience 19:298–310.

    https://doi.org/10.1523/JNEUROSCI.19-01-00298.1999

    • PubMed
    • Google Scholar
21. 1. Fabian-Fine R
    2. Meisner S
    3. Torkkeli PH
    4. Meinertzhagen IA

    (2015) Co-localization of gamma-aminobutyric acid and glutamate in neurons of the spider central nervous system

    Cell and Tissue Research 362:461–479.

    https://doi.org/10.1007/s00441-015-2241-5

    • PubMed
    • Google Scholar
22. 1. Fabian-Fine R
    2. Anderson CM
    3. Roush MA
    4. Johnson JAG
    5. Liu H
    6. French AS
    7. Torkkeli PH

    (2017) The distribution of cholinergic neurons and their co-localization with FMRFamide, in central and peripheral neurons of the spider Cupiennius salei

    Cell and Tissue Research 370:71–88.

    https://doi.org/10.1007/s00441-017-2652-6

    • PubMed
    • Google Scholar
23. Book

    1. Foelix R

    (2025)

    Spider Biology

    Springer Nature.

    • Google Scholar
24. 1. Gonzalez-Fernandez F
    2. Sherman RG

    (1984) Cardioregulatory nerves in the spider Eurypelma marxi Simon

    The Journal of Experimental Zoology 231:27–37.

    https://doi.org/10.1002/jez.1402310105

    • PubMed
    • Google Scholar
25. 1. Gorb SN
    2. Anton S
    3. Barth FG

    (1993) Central projections of cheliceral mechanoreceptors in the spider Cupiennius salei (Arachnida, Araneae)

    Journal of Morphology 217:129–136.

    https://doi.org/10.1002/jmor.1052170202

    • PubMed
    • Google Scholar
26. Software

    1. GordusLab

    (2026) Artiushin-elastix-elife, version swh:1:rev:3e853e1c7c06631d8c32e3432cfb8a9cc185a3df

    Software Heritage.

    https://archive.softwareheritage.org/swh:1:dir:464e879d437b74f72a6a3a0a4bc21d03b2d79844;origin=https://github.com/GordusLab/Artiushin-elastix-eLife;visit=swh:1:snp:ef54196478d142be93cfd3a78b156328b7fb5417;anchor=swh:1:rev:3e853e1c7c06631d8c32e3432cfb8a9cc185a3df
27. 1. Groome JR
    2. Townley MA
    3. de Tschaschell M
    4. Tillinghast EK

    (1991) Detection and isolation of proctolin-like immunoreactivity in arachnids: possible cardioregulatory role for proctolin in the orb-weaving spiders argiope and araneus

    Journal of Insect Physiology 37:9–19.

    https://doi.org/10.1016/0022-1910(91)90013-P

    • Google Scholar
28. Book

    1. Hanström B

    (1928)

    Vergleichende Anatomie Des Nervensystems Der Wirbellosen Tiere: Unter Berücksichtigung Seiner Funktion

    J. Springer.

    • Google Scholar
29. 1. Heinze S

    (2024) Variations on an ancient theme — the central complex across insects

    Current Opinion in Behavioral Sciences 57:101390.

    https://doi.org/10.1016/j.cobeha.2024.101390

    • Google Scholar
30. 1. Heuer CM
    2. Kollmann M
    3. Binzer M
    4. Schachtner J

    (2012) Neuropeptides in insect mushroom bodies

    Arthropod Structure & Development 41:199–226.

    https://doi.org/10.1016/j.asd.2012.02.005

    • Google Scholar
31. Book

    1. Hill DE

    (1975)

    The Structure of the Central Nervous System of Jumping Spiders of the Genus Phidippus

    Araneae: Salticidae.

    • Google Scholar
32. 1. Homberg U

    (2008) Evolution of the central complex in the arthropod brain with respect to the visual system

    Arthropod Structure & Development 37:347–362.

    https://doi.org/10.1016/j.asd.2008.01.008

    • Google Scholar
33. 1. Homberg U
    2. Humberg TH
    3. Seyfarth J
    4. Bode K
    5. Pérez MQ

    (2018) GABA immunostaining in the central complex of dicondylian insects

    The Journal of Comparative Neurology 526:2301–2318.

    https://doi.org/10.1002/cne.24497

    • PubMed
    • Google Scholar
34. 1. Homberg U
    2. Kirchner M
    3. Kowalewski K
    4. Pitz V
    5. Kinoshita M
    6. Kern M
    7. Seyfarth J

    (2023) Comparative morphology of serotonin-immunoreactive neurons innervating the central complex in the brain of dicondylian insects

    The Journal of Comparative Neurology 531:1482–1508.

    https://doi.org/10.1002/cne.25529

    • PubMed
    • Google Scholar
35. 1. Honkanen A
    2. Adden A
    3. da Silva Freitas J
    4. Heinze S

    (2019) The insect central complex and the neural basis of navigational strategies

    The Journal of Experimental Biology 222:jeb188854.

    https://doi.org/10.1242/jeb.188854

    • PubMed
    • Google Scholar
36. 1. Hwang H
    2. Kim H
    3. Moon M

    (2015) Immunocytochemical localization of GAD isoforms in the CNS ganglia of the cobweb spider, A chaearanea tepidariorum

    Entomological Research 45:2111.

    https://doi.org/10.1111/1748-5967.12111

    • Google Scholar
37. Book

    1. Jundi B
    2. Heinze S

    (2020)

    Three-dimensional atlases of insect brains

    In: Pelc R, Walz W, Doucette JR, editors. Neurohistology and Imaging Techniques. Springer US. pp. 73–124.

    • Google Scholar
38. 1. Kahsai L
    2. Winther AME

    (2011) Chemical neuroanatomy of the Drosophila central complex: distribution of multiple neuropeptides in relation to neurotransmitters

    The Journal of Comparative Neurology 519:290–315.

    https://doi.org/10.1002/cne.22520

    • PubMed
    • Google Scholar
39. 1. Kaiser A
    2. Hensgen R
    3. Tschirner K
    4. Beetz E
    5. Wüstenberg H
    6. Pfaff M
    7. Mota T
    8. Pfeiffer K

    (2022) A three-dimensional atlas of the honeybee central complex, associated neuropils and peptidergic layers of the central body

    The Journal of Comparative Neurology 530:2416–2438.

    https://doi.org/10.1002/cne.25339

    • PubMed
    • Google Scholar
40. 1. Klein S
    2. Staring M
    3. Murphy K
    4. Viergever MA
    5. Pluim J

    (2010) elastix: a toolbox for intensity-based medical image Registration

    IEEE Transactions on Medical Imaging 29:196–205.

    https://doi.org/10.1109/TMI.2009.2035616

    • Google Scholar
41. 1. Kollmann M
    2. Huetteroth W
    3. Schachtner J

    (2011) Brain organization in collembola (springtails)

    Arthropod Structure & Development 40:304–316.

    https://doi.org/10.1016/j.asd.2011.02.003

    • Google Scholar
42. 1. Kotsyuba E
    2. Dyachuk V

    (2021) Localization of neurons expressing choline acetyltransferase, serotonin and/or FMRFamide in the central nervous system of the decapod shore crab Hemigrapsus sanguineus

    Cell and Tissue Research 383:959–977.

    https://doi.org/10.1007/s00441-020-03309-3

    • PubMed
    • Google Scholar
43. 1. Lin C
    2. Strausfeld NJ

    (2012) Visual inputs to the mushroom body calyces of the whirligig beetle Dineutus sublineatus: modality switching in an insect

    The Journal of Comparative Neurology 520:2562–2574.

    https://doi.org/10.1002/cne.23158

    • PubMed
    • Google Scholar
44. 1. Loesel R
    2. Nässel DR
    3. Strausfeld NJ

    (2002) Common design in a unique midline neuropil in the brains of arthropods

    Arthropod Structure & Development 31:77–91.

    https://doi.org/10.1016/S1467-8039(02)00017-8

    • Google Scholar
45. 1. Loesel R
    2. Seyfarth EA
    3. Bräunig P
    4. Agricola HJ

    (2011) Neuroarchitecture of the arcuate body in the brain of the spider Cupiennius salei (Araneae, Chelicerata) revealed by allatostatin-, proctolin-, and CCAP-immunocytochemistry and its evolutionary implications

    Arthropod Structure & Development 40:210–220.

    https://doi.org/10.1016/j.asd.2011.01.002

    • Google Scholar
46. Book

    1. Long SM

    (2016)

    Spider Brain Morphology & Behavior

    University of Massachusetts Amherst.

    • Google Scholar
47. 1. Long SM

    (2021) Variations on a theme: Morphological variation in the secondary eye visual pathway across the order of Araneae

    The Journal of Comparative Neurology 529:259–280.

    https://doi.org/10.1002/cne.24945

    • PubMed
    • Google Scholar
48. 1. Martin C
    2. Jahn H
    3. Klein M
    4. Hammel JU
    5. Stevenson PA
    6. Homberg U
    7. Mayer G

    (2022) The velvet worm brain unveils homologies and evolutionary novelties across panarthropods

    BMC Biology 20:26.

    https://doi.org/10.1186/s12915-021-01196-w

    • PubMed
    • Google Scholar
49. 1. Meyer W
    2. Idel K

    (1977) The distribution of acetylcholinesterase in the central nervous system of jumping spiders and wolf spiders (Arachnida, Araneida: Salticidae et Lycosidae)

    The Journal of Comparative Neurology 173:717–744.

    https://doi.org/10.1002/cne.901730406

    • PubMed
    • Google Scholar
50. 1. Meyer W
    2. Pospiech B

    (1977) The distribution of acetylcholinesterase in the central nervous system of web-building spiders (Arachnida, Araneae)

    Histochemistry 51:201–208.

    https://doi.org/10.1007/BF00567224

    • PubMed
    • Google Scholar
51. 1. Miller J
    2. Zimin AV
    3. Gordus A

    (2022) Chromosome-level genome and the identification of sex chromosomes in Uloborus diversus

    GigaScience 12:giad002.

    https://doi.org/10.1093/gigascience/giad002

    • PubMed
    • Google Scholar
52. 1. Modi MN
    2. Shuai Y
    3. Turner GC

    (2020) The Drosophila mushroom body: from architecture to algorithm in a learning circuit

    Annual Review of Neuroscience 43:465–484.

    https://doi.org/10.1146/annurev-neuro-080317-0621333

    • PubMed
    • Google Scholar
53. 1. Moon M
    2. Tillinghast EK

    (2013) Immunoreactivity of glutamic acid decarboxylase (GAD) isoforms in the central nervous system of the barn spider, a raneus cavaticus

    Entomological Research 43:0475.

    https://doi.org/10.1111/j.1748-5967.2012.00475.x

    • Google Scholar
54. 1. Napiórkowska T
    2. Kobak J

    (2018) The allometry of the arcuate body in the postembryonic development of the giant house spider Eratigena atrica

    Invertebrate Neuroscience 18:3.

    https://doi.org/10.1007/s10158-018-0208-4

    • PubMed
    • Google Scholar
55. 1. Ott SR

    (2008) Confocal microscopy in large insect brains: zinc-formaldehyde fixation improves synapsin immunostaining and preservation of morphology in whole-mounts

    Journal of Neuroscience Methods 172:220–230.

    https://doi.org/10.1016/j.jneumeth.2008.04.031

    • PubMed
    • Google Scholar
56. 1. Park Y
    2. Kim H
    3. Kim H
    4. Moon M

    (2013) Fine structure of the CNS ganglia in the geometric spider N ephila clavata (A raneae: N ephilidae)

    Entomological Research 43:12039.

    https://doi.org/10.1111/1748-5967.12039

    • Google Scholar
57. 1. Pfeiffer K

    (2023) The neuronal building blocks of the navigational toolkit in the central complex of insects

    Current Opinion in Insect Science 55:100972.

    https://doi.org/10.1016/j.cois.2022.100972

    • PubMed
    • Google Scholar
58. 1. Rivera‐Quiroz FA
    2. Miller JA

    (2022) Micro‐CT visualization of the CNS: Performance of different contrast‐enhancing techniques for documenting the spider brain

    Journal of Comparative Neurology 530:2474–2485.

    https://doi.org/10.1002/cne.25343

    • Google Scholar
59. 1. Schmid A
    2. Duncker M

    (1993) Histamine immunoreactivity in the central nervous system of the spider Cupiennius salei

    Cell & Tissue Research 273:533–545.

    https://doi.org/10.1007/BF00333707

    • Google Scholar
60. 1. Schmid A
    2. Becherer C

    (1996) Leucokinin-like immunoreactive neurones in the central nervous system of the spider Cupiennius salei

    Cell and Tissue Research 284:143–152.

    https://doi.org/10.1007/s004410050574

    • PubMed
    • Google Scholar
61. 1. Schmid A
    2. Becherer C

    (1999) Distribution of histamine in the CNS of different spiders

    Microscopy Research and Technique 44:81–93.

    https://doi.org/10.1002/(SICI)1097-0029(19990115/01)44:2/3<81::AID-JEMT3>3.0.CO;2-O

    • Google Scholar
62. 1. Seelig JD
    2. Jayaraman V

    (2015) Neural dynamics for landmark orientation and angular path integration

    Nature 521:186–191.

    https://doi.org/10.1038/nature14446

    • PubMed
    • Google Scholar
63. 1. Senior EE
    2. Poulin HE
    3. Dobecki MG
    4. Anair BM
    5. Fabian-Fine R

    (2020) Co-expression of the neuropeptide proctolin and glutamate in the central nervous system, along mechanosensory neurons and leg muscle in Cupiennius salei

    Cell and Tissue Research 382:281–292.

    https://doi.org/10.1007/s00441-020-03217-6

    • PubMed
    • Google Scholar
64. Book

    1. Seyfarth E
    2. Hammer K
    3. Grünert U

    (1990)

    Serotonin-like Immunoreactivity in the CNS of Spiders Brain-Perception-Cognition

    Thieme.

    • Google Scholar
65. 1. Seyfarth EA
    2. Hammer K
    3. Spörhase-Eichmann U
    4. Hörner M
    5. Vullings HGB

    (1993) Octopamine immunoreactive neurons in the fused central nervous system of spiders

    Brain Research 611:197–206.

    https://doi.org/10.1016/0006-8993(93)90503-f

    • PubMed
    • Google Scholar
66. 1. Shamonin D

    (2014) Fast parallel image registration on CPU and GPU for diagnostic classification of Alzheimer’s disease

    Frontiers in Neuroinformatics 7:e0050.

    https://doi.org/10.3389/fninf.2013.00050

    • Google Scholar
67. 1. Steinhoff POM
    2. Sombke A
    3. Liedtke J
    4. Schneider JM
    5. Harzsch S
    6. Uhl G

    (2017) The synganglion of the jumping spider Marpissa muscosa (Arachnida: Salticidae): Insights from histology, immunohistochemistry and microCT analysis

    Arthropod Structure & Development 46:156–170.

    https://doi.org/10.1016/j.asd.2016.11.003

    • Google Scholar
68. 1. Steinhoff POM
    2. Harzsch S
    3. Uhl G

    (2024) Comparative neuroanatomy of the central nervous system in web‐building and cursorial hunting spiders

    Journal of Comparative Neurology 532:5554.

    https://doi.org/10.1002/cne.25554

    • Google Scholar
69. 1. Strausfeld NJ
    2. Barth FG

    (1993) Two visual systems in one brain: neuropils serving the secondary eyes of the spider Cupiennius salei

    The Journal of Comparative Neurology 328:43–62.

    https://doi.org/10.1002/cne.903280104

    • PubMed
    • Google Scholar
70. 1. Strausfeld NJ
    2. Weltzien P
    3. Barth FG

    (1993) Two visual systems in one brain: neuropils serving the principal eyes of the spider Cupiennius salei

    The Journal of Comparative Neurology 328:63–75.

    https://doi.org/10.1002/cne.903280105

    • PubMed
    • Google Scholar
71. 1. Strausfeld NJ
    2. Strausfeld CM
    3. Loesel R
    4. Rowell D
    5. Stowe S

    (2006) Arthropod phylogeny: onychophoran brain organization suggests an archaic relationship with a chelicerate stem lineage

    Proceedings. Biological Sciences 273:1857–1866.

    https://doi.org/10.1098/rspb.2006.3536

    • PubMed
    • Google Scholar
72. 1. Sukumar V
    2. Liu H
    3. Meisner S
    4. French AS
    5. Torkkeli PH

    (2018) Multiple biogenic amine receptor types modulate spider, Cupiennius salei, mechanosensory neurons

    Frontiers in Physiology 9:857.

    https://doi.org/10.3389/fphys.2018.00857

    • PubMed
    • Google Scholar
73. 1. Tarr EA
    2. Fidler BM
    3. Gee KE
    4. Anderson CM
    5. Jager AK
    6. Gallagher NM
    7. Carroll KP
    8. Fabian-Fine R

    (2019) Distribution of FMRFamide-related peptides and co-localization with glutamate in Cupiennius salei, an invertebrate model system

    Cell and Tissue Research 376:83–96.

    https://doi.org/10.1007/s00441-018-2949-0

    • PubMed
    • Google Scholar
74. 1. Thoen HH
    2. Marshall J
    3. Wolff GH
    4. Strausfeld NJ

    (2017) Insect-Like organization of the stomatopod central complex: functional and phylogenetic implications

    Frontiers in Behavioral Neuroscience 11:12.

    https://doi.org/10.3389/fnbeh.2017.00012

    • PubMed
    • Google Scholar
75. 1. Timm J
    2. Scherner M
    3. Matschke J
    4. Kern M
    5. Homberg U

    (2021) Tyrosine hydroxylase immunostaining in the central complex of dicondylian insects

    The Journal of Comparative Neurology 529:3131–3154.

    https://doi.org/10.1002/cne.25151

    • PubMed
    • Google Scholar
76. 1. Turner-Evans D
    2. Wegener S
    3. Rouault H
    4. Franconville R
    5. Wolff T
    6. Seelig JD
    7. Druckmann S
    8. Jayaraman V

    (2017) Angular velocity integration in a fly heading circuit

    eLife 6:e23496.

    https://doi.org/10.7554/eLife.23496

    • PubMed
    • Google Scholar
77. 1. Utting M
    2. Agricola HJ
    3. Sandeman R
    4. Sandeman D

    (2000) Central complex in the brain of crayfish and its possible homology with that of insects

    The Journal of Comparative Neurology 416:245–261.

    https://doi.org/10.1002/(SICI)1096-9861(20000110)416:2<245::AID-CNE9>3.0.CO;2-A

    • PubMed
    • Google Scholar
78. 1. Verasztó C
    2. Jasek S
    3. Gühmann M
    4. Bezares-Calderón LA
    5. Williams EA
    6. Shahidi R
    7. Jékely G

    (2025) Whole-body connectome of a segmented annelid larva

    eLife 13:RP97964.

    https://doi.org/10.7554/eLife.97964

    • PubMed
    • Google Scholar
79. Book

    1. Wegerhoff R
    2. Breidbach O

    (1995) Comparative aspects of the chelicerate nervous systems

    In: Breidbach O, Kutsch W, editors. The Nervous Systems of Invertebrates: An Evolutionary And1130 Comparative Approach. Birkhäuser Basel. pp. 159–179.

    https://doi.org/10.1007/978-3-0348-9219-3_9

    • Google Scholar
80. 1. Weltzien P
    2. Barth FG

    (1991) Volumetric measurements do not demonstrate that the spider brain “central body” has a special role in web building

    Journal of Morphology 208:91–98.

    https://doi.org/10.1002/jmor.1052080104

    • PubMed
    • Google Scholar
81. 1. White JG
    2. Southgate E
    3. Thomson JN
    4. Brenner S

    (1986) The structure of the nervous system of the nematode Caenorhabditis elegans

    Philosophical Transactions of the Royal Society of London. Series B, Biological Sciences 314:1–340.

    https://doi.org/10.1098/rstb.1986.0056

    • PubMed
    • Google Scholar
82. 1. Widmer A
    2. Höger U
    3. Meisner S
    4. French AS
    5. Torkkeli PH

    (2005) Spider peripheral mechanosensory neurons are directly innervated and modulated by octopaminergic efferents

    The Journal of Neuroscience 25:1588–1598.

    https://doi.org/10.1523/JNEUROSCI.4505-04.2005

    • PubMed
    • Google Scholar
83. 1. Winding M
    2. Pedigo BD
    3. Barnes CL
    4. Patsolic HG
    5. Park Y
    6. Kazimiers T
    7. Fushiki A
    8. Andrade IV
    9. Khandelwal A
    10. Valdes-Aleman J
    11. Li F
    12. Randel N
    13. Barsotti E
    14. Correia A
    15. Fetter RD
    16. Hartenstein V
    17. Priebe CE
    18. Vogelstein JT
    19. Cardona A
    20. Zlatic M

    (2023) The connectome of an insect brain

    Science 379:eadd9330.

    https://doi.org/10.1126/science.add9330

    • PubMed
    • Google Scholar
84. 1. Wolff GH
    2. Strausfeld NJ

    (2015) Genealogical correspondence of mushroom bodies across invertebrate phyla

    Current Biology 25:38–44.

    https://doi.org/10.1016/j.cub.2014.10.049

    • PubMed
    • Google Scholar
85. 1. Yim H
    2. Choe DT
    3. Bae JA
    4. Choi MK
    5. Kang HM
    6. Nguyen KCQ
    7. Ahn S
    8. Bahn SK
    9. Yang H
    10. Hall DH
    11. Kim JS
    12. Lee J

    (2024) Comparative connectomics of dauer reveals developmental plasticity

    Nature Communications 15:1546.

    https://doi.org/10.1038/s41467-024-45943-3

    • PubMed
    • Google Scholar
86. 1. Zhang M
    2. Pan X
    3. Jung W
    4. Halpern AR
    5. Eichhorn SW
    6. Lei Z
    7. Cohen L
    8. Smith KA
    9. Tasic B
    10. Yao Z
    11. Zeng H
    12. Zhuang X

    (2023) Molecularly defined and spatially resolved cell atlas of the whole mouse brain

    Nature 624:343–354.

    https://doi.org/10.1038/s41586-023-06808-9

    • PubMed
    • Google Scholar

Author details

1. Gregory Artiushin

   Department of Biology, Johns Hopkins University, Baltimore, United States

   Contribution

   Conceptualization, Resources, Data curation, Formal analysis, Funding acquisition, Investigation, Visualization, Methodology, Writing – original draft, Project administration, Writing – review and editing

   Competing interests

   No competing interests declared

   "This ORCID iD identifies the author of this article:" 0000-0003-1615-3012

2. Abel Corver

   1. Department of Biology, Lund University, Lund, Sweden
   2. Johns Hopkins Kavli Neuroscience Discovery Institute, Baltimore, United States
   3. Solomon H. Snyder Department of Neuroscience, Johns Hopkins University, Baltimore, United States

   Contribution

   Software, Formal analysis, Methodology

   Competing interests

   No competing interests declared

3. Andrew Gordus

   1. Department of Biology, Johns Hopkins University, Baltimore, United States
   2. Solomon H. Snyder Department of Neuroscience, Johns Hopkins University, Baltimore, United States

   Contribution

   Conceptualization, Supervision, Funding acquisition, Project administration, Writing – review and editing

   For correspondence

   agordus@jhu.edu

   Competing interests

   No competing interests declared

   "This ORCID iD identifies the author of this article:" 0000-0002-5550-0286

• 503

  views

• 25

  downloads

• 0

  citations

Views, downloads and citations are aggregated across all versions of this paper published by eLife.