1. Developmental Biology and Stem Cells
  2. Neuroscience
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Genetic mechanisms control the linear scaling between related cortical primary and higher order sensory areas

  1. Andreas Zembrzycki Is a corresponding author
  2. Adam M Stocker
  3. Axel Leingärtner
  4. Setsuko Sahara
  5. Shen-Ju Chou
  6. Valery Kalatsky
  7. Scott R May
  8. Michael P Stryker
  9. Dennis DM O'Leary
  1. The Salk Institute for Biological Studies, United States
  2. University of California, San Francsisco, United States
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Cite as: eLife 2015;4:e11416 doi: 10.7554/eLife.11416

Abstract

In mammals, the neocortical layout consists of few modality-specific primary sensory areas and a multitude of higher order ones. Abnormal layout of cortical areas may disrupt sensory function and behavior. Developmental genetic mechanisms specify primary areas, but mechanisms influencing higher order area properties are unknown. By exploiting gain-of and loss-of function mouse models of the transcription factor Emx2, we have generated bi-directional changes in primary visual cortex size in vivo and have used it as a model to show a novel and prominent function for genetic mechanisms regulating primary visual area size and also proportionally dictating the sizes of surrounding higher order visual areas. This finding redefines the role for intrinsic genetic mechanisms to concomitantly specify and scale primary and related higher order sensory areas in a linear fashion.

https://doi.org/10.7554/eLife.11416.001

eLife digest

The neocortex is the most recently evolved part of the human brain. It is associated with higher thought processes, including language and the processing of information from our senses. Anatomically, the neocortex is organised into different regions called ‘primary areas’ and ‘higher order areas’, and perturbations to this organisation are associated with disorders such as autism.

There are many more higher order areas than primary areas in a mammalian brain. But, while primary areas are known to be specified by developmental genes in the embryo, little is known about how the development of higher order areas is controlled. Recent findings suggested that primary areas might themselves influence the emergence of higher order areas via a series of developmental events.

Now, Zembrzycki, Stocker et al. have investigated the developmental mechanisms that organise the mouse neocortex into its different regions. The experiments involved mouse mutants that produce either too much or too little of a protein called Emx2. This protein is known to determine the size and position of the primary visual area (commonly called V1) during embryonic development. In the mutant mice with too much Emx2, the primary visual area was about 150% larger than it should be, even though the neocortex was a normal size. Zembrzycki, Stocker et al. then went on to show that the higher order areas associated with the primary visual area also grew proportionally larger in these mutant mice. The opposite was true for mice that didn’t produce Emx2 in their brains, and these mice had a much smaller primary visual area than normal mice.

Together, these findings reveal a previously unknown linear relationship between the size of the primary visual area and higher order visual areas that is controlled by the genes that pattern the neocortex during development. This and other new insights will inform future studies of the development and organization of the neocortex and our understanding of how it evolved.

https://doi.org/10.7554/eLife.11416.002

Introduction

The mouse neocortex is patterned into functionally distinct fields that include the primary sensory areas (visual, somatosensory and auditory), which receive modality-specific sensory inputs from thalamocortical axons (TCAs) originating from nuclei of the dorsal thalamus (O'Leary et al., 2013). In the cortex, the connections of TCAs establish precise topographic representations (or maps) of the sensory periphery (Krubitzer and Kaas, 2005; O'Leary et al., 2013). Primary areas are flanked by higher order sensory areas (HO), which are interconnected with them and also contain topographic maps (Felleman and Van Essen, 1991). In mammals, this evolutionarily conserved general layout of the intra-areal neural circuits is responsible for the orderly progression of sensory information, sensory perception and the integration of higher cortical functions (Felleman and Van Essen, 1991; Geschwind and Rakic, 2013; Krubitzer and Kaas, 2005; Laramée and Boire, 2014; O'Leary et al., 2013). Disrupted layouts of cortical area layouts appear to be associated with neurodevelopmental disorders including autism (Courchesne et al., 2011; Voineagu et al., 2011). Studies of cortical arealization, the mechanisms that pattern the neocortex into areas, have focused almost exclusively on the primary areas and have led to the prevailing model that genetic mechanisms intrinsic to the neocortex control arealization during early cortical development (Greig et al., 2013; Krubitzer and Kaas, 2005; O'Leary et al., 2013). For example, the graded expression of the homeodomain transcription factor Emx2 in neocortical progenitors determines the size and position of the primary visual area (V1) in mice (Bishop et al., 2000; Hamasaki et al., 2004). Although higher order areas outnumber primary areas by roughly 10-fold (Marshel et al., 2011; Wang and Burkhalter, 2007), mechanisms that specify them and define their proportions relative to primary areas have yet to be explored.

Results

To investigate the impact of altered primary area size on higher order areas, we have used the cortical visual area V1 as a model. Previous studies have shown that genetic manipulation of patterning genes, including Fgf17 and Emx2, results in altered V1 size (Cholfin and Rubenstein, 2007; Hamasaki et al., 2004). Here we have analyzed transgenic mice that overexpress Emx2 (ne-Emx2) and show area patterning defects including a V1 that is ~150% of the normal size, while retaining overall normal neocortex size (Hamasaki et al., 2004 Leingärtner et al., 2007). By revealing the targeting patterns of TCAs projecting from thalamic sensory nuclei into the cortex (Fujimiya et al., 1986), the perimeters of primary sensory areas and the border between the neocortex and entorhinal cortex (ECT) can be visualized by serotonin (5HT) staining using a single postnatal day (P) 7 tangential section of the flattened cortical hemisphere (Figure 1A). The staining shows that, in addition to the previously reported enlarged V1 (Hamasaki et al., 2004,; Leingärtner et al., 2007), the cortical tissue that is nested between V1 and the surrounding primary areas (primary somatosensory cortex: S1, auditory areas: Aud) and the ECT laterally appears qualitatively larger in ne-Emx2 brains, when compared to wildtype (wt) sections (Figure 1A). We have defined this caudal cortical territory, which lies outside of V1, S1, and the auditory areas and shows no or weak 5HT staining as a joint higher order cortical area complex and have termed it HO-5HT. The 5HTstaining revealed that this region contains the higher order visual areas surrounding V1 (Wang and Burkhalter, 2007), the retrosplenial cortex (RSC) medially, and the ventral posterior temporal cortex laterally. The accurate distribution of staining across cortical layers can only be estimated using tangential sections. However, using P7 sagittal section, we confirmed in layer 4 that the caudal 5HT-positive cortical area (corresponding to V1) and the anteriorly adjacent 5HT-negative area between V1 and S1 (corresponding to HO-5HT) appears larger in ne-Emx2 brains than in wt ones (Figure 1B). Next, we labeled TCAs projecting to V1 by filling the dLG with crystals of the lipophilic neuronal tracer DiI. On the P7 sagittal sections that were derived from five different medial to lateral levels, anterograde DiI labeling in the cortex revealed that TCAs from the dLG terminate in a smaller region in wt than in ne-Emx2 brains (Figure 1—figure supplement 1). Across genotypes, the DiI staining revealed a sharp border with adjacent cortical tissues that did not receive TCAs input from the dLG (Figure 1—figure supplement 1). This finding is consistent with the 5HT staining and indicates a well-defined border between V1 neighboring higher order areas that is anteriorly shifted in ne-Emx2 brains.

Figure 1 with 1 supplement see all
Increased V1 and higher order sensory area sizes in ne-Emx2 cortices

(A) Serotonin (5HT) staining on postnatal day (P) 7 tangential sections of the flattened cortex reveals targeting patterns of TCAs revealing primary sensory area borders and the border of the neocortex to the ECT. 5HT staining is not detectable in the region containing the retosplenial cortex and the higher order sensory areas surrounding V1 (HO-5HT). In Emx2-overexpressing brains (ne-Emx2), V1 and HO-5HT appear larger (compare dotted outlines in higher magnification images), compared to wt brains. (B) Targeting of TCAs in cortical layer 4 (L4) was revealed on P7 sagittal cortex sections by 5HT staining, whereas L4 genetic area borders were revealed by in situ hybridization for Rorb. In ne-Emx2 brains, the V1 border shifts anteriorly. Higher order areas surrounding V1 are characterized by low 5HT/Rorb staining (between arrowheads, HO-5HT and HO-Rorb), which in ne-Emx2 brains appear overall larger (compare area between arrowheads). (C) In L5, an expansion (see arrows) of corticotectal projection neurons (retrogradely labeled by DiI injections into the superior colliculus) is apparent in ne-Emx2 brains, to the expense (see dotted lines) of L5 corticospinal projection neurons (retrogradely labeled by DiI injections into the pyramidal decussation). Main axes: A: anterior; M: medial; F/M: frontal/motor cortex; S1: primary somatosensory cortex; Aud: auditory areas; V1: primary visual cortex, ECT: entorhinal cortex. 5HT, serotonin; L5, cortical layer 5; TCAs, thalamocortical axons; wt, wildtype.

https://doi.org/10.7554/eLife.11416.003

Cortical areas can also be distinguished by area-specific gene expression patterns, which overlap with anatomical area borders and shift similarly when area patterning is disrupted (O'Leary et al., 2013). For example, Rorb expression is strongly induced by thalamic input to primary areas (Jabaudon et al., 2012) like S1 and V1 but is low in areas that do not receive their major inputs from the principal thalamic sensory nuclei, such as cortical higher order areas surrounding V1 (Chou et al., 2013; Wang and Burkhalter, 2007). In situ hybridization (ISH) on sagittal sections adjacent to 5HT-stained ones revealed sharp Rorb gene expression borders between areas in layer 4. Notably in ne-Emx2 brains, the high-to-low Rorb expression border is located more anteriorly, and the area showing low Rorb expression and resembling HO-5HT (Chou et al., 2013) is larger than in wt sections (Figure 1B). This reveals that characteristic molecular markers that delineate the borders between V1 and surrounding higher order areas remain expressed at normal levels, but their sharp expression borders shift anteriorly in ne-Emx2.

Projection neurons in layer 5, which extend axons into subcortical targets, are similarly determined by a molecular code (Greig et al., 2013). We therefore predicted that the areal shifts in ne-Emx2 brains would be accompanied by corresponding changes in layer 5 output projections. We labeled two distinct types of layer 5 subcerebral projection neurons by inserting DiI crystals either into the superior colliculus, which labels corticotectal projections from V1 and HO, or else into the pyramidal decussation, which labels corticospinal projections from the frontal cortex and S1 (Greig et al., 2013; Zembrzycki et al., 2015). We found that layer 5 corticotectal projections extended more anteriorly in ne-Emx2 sagittal sections. Vice versa, the layer 5 corticospinal projections extended less posteriorly in ne-Emx2 brains. These staining patterns are consistent with an altered balance of projection neuron identity in layer 5 (Greig et al., 2013; Zembrzycki et al., 2015) and an overall expansion of visual areas, demonstrating that areal patterning changes in ne-Emx2 brains are not limited to the cortical layers that receive thalamic input. These findings complement previous reports describing Emx2 patterning functions (Bishop et al., 2000; Hamasaki et al., 2004; Leingärtner et al., 2003; Leingärtner et al., 2007) and indicate for the first time that V1 and higher order area sizes are altered concomitantly in ne-Emx2 brains at the level of area-specific connectivity and gene expression in multiple cortical layers. Taken together, our results suggest that changes in primary area size are paralleled by similar changes in higher order area size.

It is commonly assumed that areal patterning changes also alter area-specific functional neuronal properties and topographic sensory maps, but this has never been demonstrated conclusively. Therefore, to compare functional neuronal properties of an enlarged visual cortex to a normal-sized one, we have used Fourier intrinsic signal optical imaging to construct topographic visual response maps to light bars that were moved across the visual field of the retina (up and down: elevation maps; left to right: azimuth maps) (Kalatsky and Stryker, 2003). Visual responses in V1 of wt and ne-Emx2 mice produced intrinsic signal maps that were indistinguishable in strength, and the axes of azimuth and elevation were organized in the same way in all tested brains (Figure 2A), revealing that functional topographic organization of the visual cortex was intact. However, the representations of elevation and azimuth were expanded in ne-Emx2 animals, and their retinotopic maps were overall larger (elevation: 138% ± 8.7% of wt; azimuth: 143% ± 8.2% of wt). For example, the green region in the response maps representing ~20 to ~30 degrees of elevation/azimuth is clearly enlarged in ne-Emx2 brains, compared to wt brains (Figure 2A). To investigate the relationship between the location and size of the V1 functional response area and the histochemically delineated V1, as indicated by 5HT staining, multiple injections of DiI were placed lining the border of the V1 optical response map after the imaging procedure. On 5HT-stained, flattened tangential cortical sections, the DiI injection sites were found in all cases to be located near the border of the 5HT staining in V1, confirming the overall enlarged V1 perimeters in ne-Emx2 brains compared with the wt brains (Figure 2B). This shows that the 5HT-stained V1 area accurately corresponds to the intrinsic functional V1 map, suggesting that enlarged HO in ne-Emx2 brains have not acquired ectopic V1-like functional properties.

Enlarged functional V1 topographic maps in ne-Emx2 mice.

(A) Fourier intrinsic signal optical imaging reveals topographic visual response maps to light bars that were moved across the visual field of the retina (up and down: elevation maps; left to right: azimuth maps). Visual responses in V1 produced intrinsic signal maps that were indistinguishable in strength and the color-coded axes of azimuth and elevation were organized in the same way in all tested brains (wt: n = 6, ne-Emx2: n = 6). The elevation/azimuth representations were expanded in ne-Emx2 animals, revealing that overall their V1 retinotopic maps were larger. (B) 5HT staining performed on flattened tangential cortex section reveals cortical area borders including V1. On representative images (n = 6 per genotype), the red dots lining the perimeter of the 5HT-stained V1 indicates DiI injection sites that were made after the recordings adjacent to the border of the derived V1 intrinsic response maps, determined by Fourier intrinsic optical imaging. (C) Schematics depict caudal cortical sensory areas and main sensory thalamus divisions. In wt brains (n = 15), cortical dual tracer injections (red tracer (DiI) injected around V1/HO border; green tracer (DiD) into HO; injection site location (arrowheads) was identified by 5HT staining) showed retrogradely labeled red cells in the dLG and the PO, whereas green labeled cells were only present in the PO. Dotted lines show that dual tracer injections in ne-Emx2 brains (n = 17) were administered at more anterior coordinates (red tracer into V1; green tracer around the V1/HO border) compared with wt brains. In ne-Emx2 brains, retrogradely labeled red cells were apparent in the dLG, whereas green cells were present in the dLG and the PO, revealing normal thalamocortical connectivity patterns, but an anterior shifted V1/HO border in ne-Emx2 brains. 5HT, serotonin; dLG, dorsal lateral geniculate nucleus; PO, posterior thalamic nucleus; S1, primary somatosensory cortex; VP, ventroposterior nucleus; V1, primary visual cortex; wt, wildtype.

https://doi.org/10.7554/eLife.11416.005

To further characterize the shifted border between visual areas in ne-Emx2 animals, we used additional neuronal tracing approaches. Stereotypically, V1 is connected with the dLG, whereas the HO areas are wired to the posterior thalamic nucleus (PO) (Leyva-Díaz and López-Bendito, 2013; López-Bendito and Molnár, 2003). To first label axonal connections between the cortex and the thalamus in wt brains, we administered dual tracer injections into locations that approximate to HO (DiD: green dye) and another injection around the approximated border area between V1 and HO (DiI: red dye). After diffusion of the tracers, we performed 5HT staining on flattened cortex sections to identify the areas in which the injections were administered and analyzed the patterns of retrograde dye labeling on coronal sections of the thalamus. In representative cases (Figure 2C) where 5HT staining confirmed that DiI was injected at the border between V1 and HO and DiD was injected into HO, retrogradely labeled green DiD cells were apparent in the dLG and the PO. Conversely, red DiI cells were only labeled in the dLG. In ne-Emx2 brains, we administered similar dual tracer injections: DiI was targeted to V1 and a DiD injection was administered around the approximate border between V1 and HO. Due to their enlarged V1, all ne-Emx2 injections were administered at more anterior coordinates than in wt brains (compare dashed lines in Figure 2C). In representative cases (Figure 2C) where 5HT staining confirmed that the DiI injection was administered into V1 and DiD was injected around the V1/HO border, red cells were found in the dLG, whereas green-labeled cells were apparent in the PO and the dLG. The dual tracings in wt and ne-Emx2 brains are consistent with the predicted connectivity of cortical neurons around the injection sites (Leyva-Díaz and López-Bendito, 2013; López-Bendito and Molnár, 2003). Although located more anteriorly in ne-Emx2 brains, the subcortical connectivity patterns around the V1/HO borders were similar, demonstrating that these neurons show connectivity patterns that are consistent with their intrinsic areal identity and not their topographic location on the cortical sheet. Taken together, our results indicate that increased V1 size in ne-Emx2 brains is accompanied by a concomitant enlargement and anterior shift of HO.

To define the individual magnitudes of the V1 and HO size increases in ne-Emx2 brains, we next used gene expression domains as molecular markers delineating visual areas and quantified them (Figure 3). An accurate assessment of area sizes using flattened and/or sectioned cortical tissues could potentially be hampered by imperfect flattening of the tissues or by cutting artifacts. Therefore, we have used RNA in situ hybridization on intact whole brains (whole mount in situ hybridization: WMISH) at P7 for quantification purposes, which has the advantage that quantifications can be made using single images without sectioning and artifacts that may arise from such tissue processing. We first used a set of two marker genes, Unc5d and Igfbp5, whose expression delineates V1 at P7 (Chou et al., 2013). The gene expression domains on WMISH-stained brains were outlined and their sizes quantified as a measure of V1 area size. The mean value of wt brains was defined as 100% and the area size percentages of ne-Emx2 brains displayed accordingly as ‘percent of wt’ (Figure 3A). V1 gene expression domains of both markers were larger in ne-Emx2 brains (Unc5d-V1: 148% ± 6.1%, p = 0.0003; Igfbp5-V1: 148 ± 4.5%, p < 0.0001). The magnitude of the increased in V1 size labeled genetically in ne-Emx2 brains is comparable to V1 area measurements derived from 5HT-stained P7 flattened cortical sections (Figure 3A: 5HT-V1: 142.1% ± 3.1%, p < 0.0001), indicating that molecular markers on whole brains reliably delineate V1 and can therefore be used to quantify and compare area sizes between samples and mouse lines.

Figure 3 with 2 supplements see all
Proportionally increased V1 and HO sizes in ne-Emx2 cortices.

(A) Schematic shows sensory area outlines in the caudal neocortex (12). WMISH with the molecular V1 marker genes Unc5d (wt: n = 5, ne-Emx2: n = 6) and Igfbp5 (wt: n = 11, ne-Emx2: n = 6) at P7 highlights increased V1 size in ne-Emx2 brains using whole un-sectioned brains. Quantification of V1 size using 5HT-stained P7 flattened cortical sections similarly reveals larger V1 sizes in ne-Emx2 brains (n = 11), compared to wt brains (n = 13). (B) WMISH for molecular markers that label both V1 and HO (dotted outlines: Cdh8, Lmo4: high expression in HO, lower in V1; for each probe and genotype n = 6) reveal that V1 as well as HO sizes in ne-Emx2 are larger compared with wt brains. Cdh8 is not expressed around the anteromedial edge of V1 (arrowheads). Quantifications in Figures 3 and 4 show mean values as percent of wt, error bars indicate standard error of the mean; asterisks highlight statistical significance according to unpaired to t-test. 5HT, serotonin; S1, primary somatosensory cortex; WMISH, whole mount in situ hybridization; V1, primary visual cortex; wt, wildtype.

https://doi.org/10.7554/eLife.11416.006

We next have used additional markers to quantify higher order area sizes (Figure 3B). Previous studies have parsed higher order visual areas using neuroanatomical tracers (Wang and Burkhalter, 2007) (see also schematic in Figure 3) and have revealed genes that are expressed at different levels in V1 and higher order visual areas (Chou et al., 2013). For example, Cdh8 and Lmo4 expression is higher in the area surrounding V1, where higher order visual areas are located (Chou et al., 2013; Marshel et al., 2011; Wang and Burkhalter, 2007). The domains of high Cdh8 and Lmo4 expression appear to label higher order visual areas uniformly (Chou et al., 2013), without revealing subdivisions between them (compare schematic in Figure 3 showing approximate location and outline of higher order visual areas as identified by Wang and Burkhalter, 2007 to Cdh8 and Lmo4 gene expression domains around V1). On P7 WMISH-stained wt brains, we quantified the V1 and HO sizes in the medial cortex on the basis of low and high gene expression domains (see dotted lines in Figure 3, Figure 3—figure supplement 2). Anatomically, these gene expression domains surrounding V1, which show much stronger staining compared with V1, extend anteriorly up to the S1 border, laterally to the border of the auditory areas and the ECT and medially up to the border to the RSC, respectively (Figure 3—figure supplements 1,2). Hence, compared with the above-mentioned HO complex that was identified using 5HT staining (Figure 1), the higher order area complex labeled by Cdh8 and Lmo4 relates to a smaller cortical region that more closely relates to higher order visual areas, but excludes the RSC. The size (Cdh8-V1: 145.3 ± 7.4%, p < 0.0001; Lmo4-V1: 146.6% ± 6.7%, p = 0.0005) and shape of the gene expression domains in V1 were similar in Cdh8- and Lmo4-stained brains. Similarly, the gene expression domains nested around V1 largely overlapped between the two probes. The only apparent difference between them is around the anteromedial edge of the higher order visual areas (Wang and Burkhalter, 2007), where Cdh8 is expressed at much lower levels compared to more lateral regions around V1 across genotypes (arrowheads in Figure 3B). The wt values of the measurements were again defined as 100%. The overall shapes of the two HO marker gene domains were similar and the sizes larger in ne-Emx2 brains compared with wt brains (Cdh8- HO: 145.7 ± 6.4%, p = 0.0015; Lmo4- HO: 144.9 ± 3.8%, p = 0.00157). The analysis of different area-specific sets of marker genes, either showing unique expression in V1, or discernable expression levels between visual areas, revealed an increase in visually-related HO in ne-Emx2 brains that was proportionate to the V1 size increase. The extrastriate areas that we have identified on the basis of 5HT staining (Figure 1) included the RSC, which is not a higher order visual area (Garrett et al., 2014; Marshel et al., 2011; Vann et al., 2009; Wang and Burkhalter, 2007) raising the possibility that only related primary and higher order areas (e.g. vision) could scale proportionately. To test this possibility, we have used WMISH of Lypd1 on P7 wt and ne-Emx2 brains as a specific marker labeling the caudomedial cortex, where the RSC is located (Figure 3—figure supplement 1). We found that the specific Lypd1 gene expression domain in the caudomedial cortex is significantly enlarged in ne-Emx2 brains (114.3 ± 5.2%, p = 0.0225, n = 4), compared with wt brains. This size increase is not proportionate to the size increases of V1 and the higher order visual area complex labeled by Cdh8 and Lmo4 in ne-Emx2 brains (Figure 3) suggesting that increased V1 size is specifically accompanied by a proportionate size increase of related higher order visual areas.

To test if related HO size matches V1 size only when it is larger than normal, or if primary area size bi-directionally is accompanied by according scaling of related higher order areas, we next analyzed HO sizes excluding the RSC in brains with a smaller than normal V1 (Figure 4). Constitutive Emx2 mutant mice have an overall smaller brain and visual cortex, but homozygous mutants die perinatally (Bishop et al., 2000), preventing the analysis of cortical areas, which arise at later stages. To overcome this limitation, we generated a novel mouse line with floxed Emx2 alleles (Figure 4—figure supplement 1), allowing conditional inactivation of Emx2. We crossed Emx2 floxed mice with Emx1-IRES-Cre expressing mice (Gorski et al., 2002) to generate conditional, cortex-specific Emx2 mutant mice. These cKO mice are viable, fertile and have an anatomically normal neocortex (Figure 4—figure supplement 2). Confirming the prediction that reduced Emx2 expression levels in cortical progenitors would lead to smaller visual areas, cKO brains show areal patterning changes (Figure 4—figure supplement 3) that are similar to those previously reported in heterozygous Emx2 mutant brains (e.g. larger frontal cortex) (Hamasaki et al., 2004), but are opposite to those apparent in ne-Emx2 brains (e.g. smaller frontal cortex) (Hamasaki et al., 2004; Leingärtner et al., 2007). As in ne-Emx2 brains, V1 in cKO brains was characterized using 5HT staining and DiI injections into the dLG (Figure 4—figure supplement 4), revealing that V1 in cKO is greatly reduced in size.

Figure 4 with 4 supplements see all
Proportionally decreased V1 and HO sizes in cKO cortices.

WMISH for V1 (A: Unc5d; wt: n = 5, ne-Emx2: n = 6, Igfbp5; wt: n = 11, ne-Emx2: n = 8) or V1 and HO marker genes (B: Cdh8; wt: n = 6, ne-Emx2: n = 6; Lmo4; wt: n = 5, ne-Emx2: n = 5) at P7 conversely reveals decreased sizes (~70% of wt size) of V1 and HO in brains that were derived from Emx1-IRES-Cre-mediated cortex-specific conditional Emx2 mutant brains (cKO), compared with wt brains. Quantification of V1 size using 5HT staining on P7 flattened cortical sections (A: wt: n = 15, cKO: n = 10) reveals similar reductions of V1 size in cKO brains. (C) The ratio between quantified V1 and HO sizes derived from WMISH-stained brains with decreased (cKO), normal (wt), and increased (ne-Emx2) V1 sizes demonstrates linear scaling of HO size in response to bi-directional changes of V1 size. S1, primary somatosensory cortex; V1, primary visual cortex; WMISH, whole mount in situ hybridization; wt, wildtype.

https://doi.org/10.7554/eLife.11416.009

To complement the quantification of V1 and its related HO sizes in ne-Emx2 brains, we next have used cKO brains to perform WMISH with both sets of marker genes noted above (Figure 3) and measured their sizes (Figure 4). Measurements of the molecular V1 marker domains (Unc5d-V1: 67.1 ± 2.6%, p = 0.00029; Igfbp5-V1: 68 ± 3.3%, p = 0.00017), as well as the V1 expression domains of Cdh8 (67.7 ± 4.5%, p < 0.0001) and Lmo4 (68.6 ± 0.8%, p < 0.0001) revealed that the molecularly defined V1 in cKO was smaller than in wt brains. These reductions matched the reduced V1 in 5HT-stained flattened cortex sections (Figure 4A: 5HT-V1: 68.5 ± 3%, p < 0.0001). Subsequent quantification of the gene expression domains of Cdh8 and Lmo4 surrounding V1 (Figure 4B: Cdh8- HO: 66.8 ± 2.9%, p < 0.0001; Lmo4- HO: 69.3 ± 2.4%, p < 0.0001) revealed that the cortical region that contains visually-related HO was also reduced in cKO brains to a degree proportional to the reduction in V1 size. These data demonstrate that when V1 size is reduced, related HO size is reduced to a similar extent.

In order to reveal a correlation between primary and related higher order area size between brains with larger and smaller than normal visual areas, we calculated the ratios between the genetically defined V1 and related HO sizes (Figure 4C). Ranging over an ~80% variation of the normal V1 size, Figure 4C reveals that related HO size is bi-directionally altered in a linear fashion (Cdh8 regression: y = −0.0071x + 1.7463; Lmo4 regression: y = 0.0208 + 1.7463). Taken together, our results are consistent with a proportional scaling relationship between the size of primary and related higher order visual areas: The size of V1 is determined by the activity of transcription factors including Emx2 during development, and this mechanism likewise controls the linear matching of the proportions of higher order visual areas in the mouse neocortex.

Discussion

The present findings address the mechanisms that specify and regulate the size of higher order sensory areas, an issue that has been largely neglected. They reveal a novel, prominent role for intrinsic genetic information in this process. Genetically altering the size of V1 over a range of ~80% of its normal size using a Emx2 gain-of function mouse line and a novel conditional Emx2 loss-of function mouse line showed that the specification of both primary and related higher order cortical areas during development was linearly scaled by driving the unique properties that characterize both, V1 and higher order visual areas.

Regardless of whether V1 was larger or smaller than in wt mice, related HO exhibited normal cytoarchitecture, genetic profiles, functional properties, and characteristic patterns of connectivity that resulted in an overall uniformly altered ‘visual cortical field’ in the occipital cortex that remained accurately and proportionally subdivided into V1 and higher order visual areas. This demonstrates that Emx2 (and perhaps additional intrinsic area patterning regulators) specify a ‘sensory cortical field’ that includes primary and higher order areas and a defined border between them. This model of cortical area patterning is not consistent with the possibility that the core properties of primary and higher order areas are specified sequentially or through parallel genetic mechanisms.

Our results further reveal that higher order areas do not have a fixed size. Rather their relative size is flexible. By using mouse models with bi-directional changes of V1 size as a model, our study revealed that higher order areas scale linearly together with their related primary sensory areas, This observation is important for at least two reasons: (i) it re-emphasizes a sequential model of primary sensory area formation that likewise influences the properties of related higher order areas. In this model, cortical intrinsic mechanisms specify all generic primary and higher order visual cortex properties during early development. Much later during postnatal development, geniculocortical input is needed to terminally differentiate the genetic distinctions between V1 and HO (Chou et al., 2013; Vue et al., 2013). (ii) It contradicts the hypothesis that cortical structure/function evolution mainly is driven by a disproportionate increase in the size of related higher order areas relative to primary areas. To the contrary, our results show that primary and related higher order areas remain proportionate when primary area size is altered through genetic mechanisms, suggesting that an increase in the complexity of connections and micro-circuits among higher order cortical processing centers likely accounts for gains in cortical functions that are characteristic for gyrencephalic mammals with larger cortical surface areas, compared to simpler lissencephalic mammals. In summary, the newly discovered linear scaling relationship between primary and related higher order areas has major implications for the basic understanding of the development and organization of the neocortical bauplan and its evolution and variability in normal and affected conditions.

Materials and methods

Mouse lines and conditional Emx2 gene targeting

All experiments were approved and conducted following the guidelines of the Institutional Animal Care and Use Committee at the Salk Institute and were in full compliance with the guidelines of the National Institutes of Health for the care and use of laboratory animals. When mice were mated, the morning of the identified vaginal plug was designated as E0.5. The morning on which pups were born was designated P 0.5. Transgenic mice overexpressing Emx2 under the Nestin promoter (ne-Emx2) were previously described (Hamasaki et al., 2004). For generating Emx2 floxed mice (Emx2fl/fl), gene targeting was carried out using homologous recombination in embryonic stem cells. A targeting construct was designed in which the 5’ loxP site was upstream of the Emx2 transcriptional start site and the 3’ loxP site downstream of Exon 2, followed by a FRT-site-flanked PGK-Neo cassette, Figure 4—figure supplement 1). Targeted embryonic stem cell clones were screened by Southern blot with probes A, B, and C and by PCR to identify Emx2floxed-neo/+ clones (Figure 4—figure supplement 1). Positive clones were injected into C57BL/6J blastocysts at the Salk Transgenic Core Facility and chimeras were mated to C57BL/6J females to obtain germline transmission. Heterozygous mice were mated with mice expressing FLPe (Rodríguez et al., 2000) to remove the neo cassette and then mated to obtain homozygous Emx2fl/fl mice. Cortex specific deletion of Emx2 (cKO) was obtained by crossing Emx2fl/fl mice with Emx1-IRES-Cre mice (Gorski et al., 2002). Specificity of Emx1-IRES-Cre-mediated deletion of Emx2 floxed alleles was analyzed by WMISH (described below) staining using a full-length Emx2 antisense RNA probe on E11 embryos. Genotyping was performed using primers for Emx2 floxed alleles (Emx2 forward: GAC-TCC-TTT-CCC-AAA-TAA-CCC-C, Emx2 reverse: GTA-AGC-GGG-CGG-GGA-CTG-GTT-C) and for the Cre recombinase (cre forward: GCT-AAA-CAT-GCT-TCA-TCG-TCG-G, cre reverse: GAT-CTC-CGG-TAT-TGA-AAC-TCC-AGC), and the ne-Emx2 transgene (nestin forward: TCA-ACC-CCT-AAA-AGC-TCC, Emx2 reverse: GGA-CGG-AGA-GAA-GGC-GGT).

In situ hybridization, immunostainings, and tangential cortical sections

Tissues were dissected, washed in phosphate-buffered saline (PBS), fixed overnight in 4% phosphate-buffered paraformaldehyde (PFA), washed in PBS, and cryopreserved in 30% sucrose in PBS. Postnatal brains were perfused with PFA, postfixed overnight in PFA, washed with PBS, and cryopreserved in 30% sucrose in PBS. Tissues were embedded in Tissue-Tek OCT (Sakura Finetek , Japan) and sectioned on a cryostat (Leica, Germany). Antisense RNA probes were labeled using a DIG-RNA labeling kit (Roche, Switzerland). ISH on 18-μm cryostat sections and WMISH using P7 brains were carried out as previously described (Chou et al., 2013; Hamasaki et al., 2004; Zembrzycki et al., 2007; Zembrzycki et al., 2015). For tangential cortical sections, cortical hemispheres were dissected, flattened, postfixed between slide glasses, and then cryoprotected. Tangential sections were cut into 40-μm slices from flattened cortical hemispheres with a sliding microtome and then they were immunostained for Serotonin (5HT, ImmunoStar, Hudson, WI). Immunostaining was developed using the diaminobenzidine colorimetric reaction and the Vectastain kit (Vector, Burlingame, CA). For Nissl staining, sections were stained with 0.5% cresyl violet and then dehydrated with graded alcohols.

Axon tracings

Lipophilic tracers DiI and DiD (all from Molecular Probes, Eugene, OR) were used to label corticothalamic-, thalamocortical-, corticotectal-, and corticospinal projections. For each experiment 4–6 brains with comparable tracer injection sites were cut and used for further data analysis, representative example images are shown in the figures. Analysis of thalamocortical axons by thalamic DiI injections (Figure 1B): P7 brains were fixed in 4% PFA, hemisected, and a coronal cut between the diencephalon and mesencephalon was made in order to expose thalamic nuclei at the section surface and DiI crystals were implanted to cover the dorsal lateral geniculate nucleus (dLG). After incubation for 1 to 2 months at 30°C to 60°C, preparations were sectioned sagittally on a vibratome (Leica). Sections were counterstained with DAPI (Vector) and analyzed under a fluorescence microscope to determine the tangential distribution of labeled thalamocortical axons in the neocortex.

Analysis of layer 5 subcerebral projection neurons (Figure 1C): Corticospinal neurons in cortical layer 5 were retrogradely labeled by inserting DiI crystals into the pyramidal decussation in 4% PFA fixed brains. Layer 5 corticotectal neurons were labeled in 4% PFA fixed brains by implantation of small DiI crystals into the upper layers of superior colliculus. Brains were incubated at 37°C for 2–3 months before 100 μm sagittal vibratome sections were cut and analyzed under fluorescent light. Analysis of area-specific thalamocortical and corticothalamic connectivity of caudal cortex (Figure 2): P7 pups were anesthetized by hypothermia and a small area of skull was removed to expose the cortical surface. DiI crystals and a small piece of DiD were implanted into cortical locations around V1 and the V1/HO border. After 1 day of survival, brains were removed after 4% PFA perfusion and their cortices and thalami dissected. Cortices were then flattened and stained for 5-HT to reveal primary sensory areas relative to the dye injection sites. The thalami were preserved sectioned coronally, stained with DAPI and DiI/DiD labeled cells analyzed under a fluorescence microscope.

Statistical analysis, area measurements, and intrinsic signal optical imaging

Data collection and analyses were performed blind to genotype and the conditions of the experiments, data were collected and processed randomly, and no data points were excluded. No statistical methods were used to predetermine sample sizes, but our sample sizes were similar to those reported in previous publications (for example, (Chou et al., 2013; Zembrzycki et al., 2013). Data met the assumptions of the statistical tests used, and the data distribution was assumed to be normal but was not formally tested. Statistics were calculated with Microsoft Excel. Quantifications show mean values of the tested groups and are displayed as a percentage of the wt group. Quantified sample sizes (number of brains: n) are indicated in the figure legends. The examples shown in each figure are representative and were reproducible for each set of experiments. Individual experiments were successfully repeated at least three times using different litters.

Area size measurements on 5HT stained sections and statistical analysis was performed as previously described (Leingärtner et al., 2007; Zembrzycki et al., 2015; 2013). To quantify molecular V1 and HO sizes, gene expression domains were quantified on single images of WMISH-stained brains (examples of measured area outlines are shown as dashed lines in Figures 3,4) using ImageJ (Rasband 1997−2013). Derived wt mean values were defined as 100% and values of the other mouse lines calculated accordingly. Statistical significance was determined using unpaired two-tailed t test, p values < 0.05 (indicated as *) were considered as statistically significant. Variance is indicated in the main text sections reflecting standard error of the mean. Intrinsic signal optical imaging was performed as previously described (Kalatsky and Stryker, 2003). To determine the spatial relationship between V1 functional maps and V1, defined histochemically by 5-HT staining, animals were imaged to determine the V1 map, and after completion, small DiI injections were made outside of the functional map perimeters. Animals were then perfused with 4% PFA, cortices dissected, flattened, sectioned tangentially, and stained for 5-HT.

References

  1. 1
  2. 2
    Patterning of frontal cortex subdivisions by Fgf17
    1. JA Cholfin
    2. JL Rubenstein
    (2007)
    Proceedings of the National Academy of Sciences of the United States of America 104:7652–7657.
    https://doi.org/10.1073/pnas.0702225104
  3. 3
  4. 4
  5. 5
  6. 6
  7. 7
  8. 8
  9. 9
    Cortical excitatory neurons and glia, but not GABAergic neurons, are produced in the Emx1-expressing lineage
    1. JA Gorski
    2. T Talley
    3. M Qiu
    4. L Puelles
    5. JL Rubenstein
    6. KR Jones
    (2002)
    The Journal of Neuroscience : The Official Journal of the Society for Neuroscience 22:6309–6314.
  10. 10
  11. 11
  12. 12
  13. 13
    New paradigm for optical imaging: temporally encoded maps of intrinsic signal
    1. VA Kalatsky
    2. MP Stryker
    (2003)
    Neuron 38:529–545.
  14. 14
  15. 15
  16. 16
  17. 17
  18. 18
  19. 19
  20. 20
  21. 21
    Area Patterning of the Mammalian Cortex
    1. DDM O'Leary
    2. AM Stocker
    3. A Zembrzycki
    (2013)
    In: DDM O'Leary, AM Stocker, A Zembrzycki, Rubenstein John, Rakic Pasko, editors. Patterning and Cell Type Specification in the Developing Cns and Pns. Oxford: Elsevier. pp. 61–85.
    https://doi.org/10.1016/B978-0-12-397265-1.00021-6
  22. 22
  23. 23
  24. 24
  25. 25
  26. 26
  27. 27
    Area map of mouse visual cortex
    1. Q Wang
    2. A Burkhalter
    (2007)
    The Journal of Comparative Neurology 502:339–357.
    https://doi.org/10.1002/cne.21286
  28. 28
  29. 29
  30. 30

Decision letter

  1. Moses V Chao
    Reviewing Editor; New York University School of Medicine, United States

eLife posts the editorial decision letter and author response on a selection of the published articles (subject to the approval of the authors). An edited version of the letter sent to the authors after peer review is shown, indicating the substantive concerns or comments; minor concerns are not usually shown. Reviewers have the opportunity to discuss the decision before the letter is sent (see review process). Similarly, the author response typically shows only responses to the major concerns raised by the reviewers.

Thank you for submitting your work entitled "Genetic mechanisms control the linear scaling between cortical primary and higher order sensory areas" for consideration by eLife. Your article has been reviewed by three peer reviewers, and the evaluation has been overseen by a Reviewing Editor and David Van Essen as the Senior Editor.

The reviewers have discussed the reviews with one another and the Reviewing editor has drafted this decision to help you prepare a revised submission.

The three reviewers found this study of visual cortical size regulation to be cleverly designed and of interest. The manuscript revisits earlier studies on the regulation of higher visual cortex specification and provides new insights into the genetic basis of cortical development.

Several major issues were brought up by the reviewers. One concern is the way the topographic locations of VI and how the parcellations of VHO were defined. It was felt the effects were not conclusive due to potentially imperfect flat-mounting of cortex and inconsistent delineation of V1 and higher order cortex. Another related reservation is the lack of evidence for reliable areal parcellation. Although the experimental design of manipulating the size of V1 is clever and elegant, without reliable parcellation of the cortex, the results fall short of being convincing to demonstrate that the size of V1 is scaled and that there is a clear cause and effect between a change in the size of V1 and the size of neighboring higher order areas.

The referees raised several specific issues that were deemed essential. They requested that the revised manuscript demonstrate that: (1) the size of V1 in control and experimental brains was determined in identical fashion; (2) that the measurements of the borders of VHO are clearly outlined in control and experimental brains; (3) The borders of the extrastriate visual cortex need to be better defined. Does VHO exclude (or include) retrosplenial cortex; and (4) the lateral border of VHO relative to primary auditory cortex, ventral posterior temporal cortex and rhinal fissure needs to be clearly defined. More detailed comments directly related to these concerns are elaborated below with recommendations for improvement:

1) The VHO in the EMX2 case includes retrosplenial cortex and extends deeper into temporal cortex than in WT animals. This raises the questions how the borders of VHO were defined and what types of criteria for flattening the cortex were used to compare the sizes of V1 and VHO. There is strong evidence that VHO extends far beyond extrastriate visual cortex. Thus, one option is to drop the "V" and call the region "HO".

2) Figure 1B. Labeling of the thalamocortical input is challenging. It is difficult to rule out that labeling patterns are produced by injections at particular topographic locations. The experiment should indicate the LGN injection site and the projections in flatmounted cortex, superimposed onto 5HT labeled V1 in the same mouse.

3) Figure 1C. The analysis could be strengthened by 5HT staining of parallel sections showing S1 and V1.

4) Figure 2A. A more compelling demonstration of the change in the size of V1 and VHO would be to mark reference points in the topographic map, stain the same flattened cortex for 5HT and superimpose the images. Further, it would strengthen the interpretation of the data if the maps included information about the topographic representations in areas of the lateral extrastriate cortex.

5) Figure 2B. The cartoon of the mouse thalamus and the labels are inaccurate for the DiI and DiD marked projections. The DiI and DiD injections into WT and ne-Emx2 mice are targeted to different topographic locations of V1 and areas within VHO. These discrepancies weaken the conclusion that the corticothalamic projections are comparable.

6) Figure 3B. The borders of VHO in WT and ne-Emx2 mice are different. Again VHO is ill-defined and its borders are not consistently outlined. For example, the Cad8 (WT) stained region around the tip and continuing along the medial border of V1 is much larger than indicated by the stippled line. In addition, V1 in ne-Emx2 mice has an unusual shape with an unexpected "tipped nose", indicating that the region enclosed by the stippled line may be larger than V1. The flattening technique can contribute to an overestimation of the size differences reported in the study.

7) Figure 4B, similar to Figure 3B. Cad8 (WT) and Lmo4 (WT) include weakly stained regions in medial extrastriate cortex, which are not included in cKO mice.

[Editors' note: further revisions were requested prior to acceptance, as described below.]

Thank you for resubmitting your work entitled "Genetic mechanisms control the linear scaling between cortical primary and higher order sensory areas" for further consideration at eLife. Your revised article has been evaluated by David Van Essen (Senior Editor), a Reviewing Editor, and two reviewers. The manuscript has been improved but there are some remaining issues that need to be addressed before acceptance, as outlined below:

The two prior reviewers indicated the paper has been improved by additional 5HT data to address the location of V1 with other cortical regions and with the additional analysis of the borders of adjacent areas by gene markers, Lmo4 and Cad8. The modifications and explanations in the revision help to clarify the questions that were previously raised. The new images that are provided are impressive and strengthen the study.

However, there is still a reservation about how V1 and HO are identified and defined. Re-evaluation of the figures indicates that the boundaries of HO differ from one figure to another. Specifically, in Figure 1A, the border of HO runs from the anterior retrosplenial cortex along the border of auditory cortex and ends near the rhinal fissure. But in Figure 3A, HO does not extend to the border of auditory cortex or include the retrosplenial cortex. Hence, there are two different criteria used to define HO in the manuscript. Furthermore, Cdh8 in WT and ne-Emx2 extends more medially than indicated in Figure 3B and also in Figure 4B, leading to a discrepancy in the analysis.

It is strongly recommended that a re-analysis of the data take place with a critical eye to establish a more consistent segmentation scheme for defining the detailed borders of HO. One suggestion is to measure HO as it is presented in Figure 1A and to exclude retrosplenial cortex. The resulting region would represent a large amount of visually-related territory.

The previous review indicated there was not enough evidence to conclude that V1 size controls the sizes of surrounding cortical areas. Therefore, the Abstract should be modified to emphasize that genetic factors intrinsic to the cortex proportionately dictate the size of primary visual cortex and surrounding higher order areas.

Reviewer #1:

The authors have very largely addressed the suggestions of the previous review, and their paper now seems ready for publication.

One remaining point: The Abstract needs to be slightly rewritten to bring it into line with the rest of the paper.

In the Abstract it is stated: "we have generated bi-directional changes in primary visual cortex size in vivo and have used it as a model to show a novel and prominent function for genetically determined primary visual area size, which also proportionally dictates the sizes of higher order visual areas." It seems (again) that the authors want to say that V1 size controls the size of surrounding higher order areas. Previous review suggested there was not enough evidence for such a conclusion, and pointed to an alternate interpretation: genetic factors intrinsic to cortex control the size of both V1 and HO areas in coordination. The cover letter for the revised MS and the new Discussion section appear to agree with the latter conclusion.

Reviewer #3:

This is a reply to comments about how primary visual cortex and the surrounding higher order cortex were delineated and the relative size of these cortical parcels were measured and compared. The detailed reply indicates that the authors clearly understood the critique and made a number of revisions in the figures and the text that significantly improved the manuscript. Although V1 and the organization of surrounding cortex are beautifully illustrated in several newly added images, there are remaining concerns that the revision failed to analyze the outstanding material in a consistent fashion. This impression is based on the inexplicable discrepancy of the definition of HO shown in Figure 1A and Figure 3A, which indicates that the authors are unsure how to subdivide the real estate. In Figure 1A it appears that the border of HO runs from the anterior tip of retrosplenial cortex along the posterior border of S1, continues along the border of auditory cortex and ends by intersecting the rhinal fissure. This does not fit the scheme shown in Figure 3A, in which HO does not extend to the border of auditory cortex and does not include retrosplenial cortex. If I understand this correctly, HO in Figure 3A is clearly smaller than in Figure 1A and does not include ventral posterior temporal cortex nor does it include retrosplenial cortex. In fact, HO Figure 3A is what was measured in Figure 3 and Figure 4. Maybe these discrepancies were simply an oversight which can be easily corrected. But the lapse heightened my concerns that the size of HO was inconsistently assessed (e.g. in Figure 3B the border of Cdh8-expression in wt is much more medial than indicated and differs from the border in ne-Emx2) and that the substance of the paper is less solid than suggested by the graphs shown in Figure 3B and Figure 4B. I recommend re-analyzing the material with a more critical eye on the detailed borders of HO. I am sure that the authors are acutely aware that details matter a lot more in small than large brains.

[Editors' note: further revisions were requested prior to acceptance, as described below.]

Thank you for resubmitting your work entitled "Genetic mechanisms control the linear scaling between related cortical primary and higher order sensory areas" for further consideration at eLife. Your latest revision has been favorably evaluated by David Van Essen (Senior Editor), a Reviewing Editor, and the two prior reviewers. There are some remaining issues that need to be addressed before acceptance, as outlined below:

Reviewer #3:

I am encouraged by the revision in paragraph seven, Results, which now more clearly defines the borders of HO. Several recommendations are made to clarify the following points.

1) To resolve the conflict between the measured HO (Figure 3, Figure 4) and HO defined by 5HT staining (or the virtual absence of 5HT expression) in Figure 1, I suggest renaming it HO-5HT to indicate the conditions under which the observation was made and unequivocally state that HO and HO-5HT are not the same.

2) HO includes at a minimum visually dominated as well as higher order auditory areas. I therefore recommend revising the sentence (Abstract) to: "…surrounding higher order sensory areas". I further recommend that similar revisions be made throughout the text to make it absolutely clear that the cortex measured here is more extensive than extrastriate visual cortex.

3) I recommend revising the border of HO (wt/Cdh8) in Figure 3B. It is clearly evident in the image that the Cdh8 expression includes a wedge medial to V1 which extends to the tip of the arrow head. A similar adjustment of the medial border is necessary for ne-Emx2 Cdh8 (Figure 3B). The same revisions are necessary in Figure 4B in which the medial borders of Cdh8 expression in wt and cKO are more medial than indicated.

https://doi.org/10.7554/eLife.11416.014

Author response

The three reviewers found this study of visual cortical size regulation to be cleverly designed and of interest. The manuscript revisits earlier studies on the regulation of higher visual cortex specification and provides new insights into the genetic basis of cortical development.

Several major issues were brought up by the reviewers. One concern is the way the topographic locations of VI and how the parcellations of VHO were defined.

To address the concern raised by the reviewers regarding how the topographic locations of V1 were defined, we have revised the manuscript to better explain our approach and results. In general in Figure 1 and Figure 2, we introduce V1 along with the other cortical areas on the basis of 5HT staining that labels thalamocortical input, with area-specific marker genes like Rorb and by using functional imaging. All of these approaches have been used in many studies from many different labs over many years with consistent results. We feel, for example, that there is no doubt that the triangular-shaped 5HT domain in the caudal cortex (Figure 1A) is V1. Similarly, we do not hesitate to state that the functional imaging maps (Figure 2A) reveal V1 and its topography. To address the reviewer’s concern, we have added 5HT images showing the entire cortical hemisphere into Figure 1A to better illustrate the location of V1 and all the other apparent cortical areas and regions on the entire cortical sheet. This now demonstrates much more clearly where, for example, the border between neocortex and entorhinal cortex is located and that 5HT staining is not apparent in the retrosplenial cortex. On the adjacent higher magnification images, we then demonstrate that extrastriate areas including retrosplenial cortex appear enlarged in ne-Emx2 brains qualitatively. We have revised the text accordingly to implement this information and to better define HO (paragraph one, Results).

In regard to the parcellation of HO in Figure 1, we have revised the text accordingly. As it now is more clearly demonstrated on the low magnification images, HO areas are now defined as follows: “We have defined this caudal cortical territory, which lies outside of V1, S1, and Aud and shows no or weak 5HT staining as a joint higher order cortical area complex (HO). Based on the 5HT staining, this region contains the higher order visual areas surrounding V1 (Wang and Burkhalter 2007), the retrosplenial cortex medially, as well as the ventral posterior temporal cortex laterally.”

In this study, we specifically avoid making claims about parcellation within higher order visual areas, since to our knowledge no marker genes exist that would allow to conclusively distinguish the at least 7-9 (Marshel et al. 2011, Wang and Burkhalter 2007) different higher order visual areas from one another.

It was felt the effects were not conclusive due to potentially imperfect flat-mounting of cortex and inconsistent delineation of V1 and higher order cortex.

We agree with the reviewers’ comments that flat-mounting potentially can have an effect on the measurement or interpretation of the data. However, we do not think that such artifacts have a major influence on the conclusions of our study for several reasons. Except for Figure 1A, we mostly use sagittal sections or un-sectioned whole-mount brains (Figure 3 and Figure 4). This is especially true for the quantifications of V1 and HO that were exclusively derived from images of unsectioned whole brains. In all other instances where we use flattened cortical sections, we only make qualitative claims about the relative area proportions (e.g. V1 is bigger or smaller). We emphasize that all specimens were processed identically which suggests that possible artifacts would affect control and experimental cases equally, still allowing comparisons between brains. For example, it is very common that the staining of the auditory areas using 5HT staining on flattened sections is a bit fuzzy, while area borders more medially in the cortex are almost always very sharp (e.g. barrels). As we all know the auditory area borders are in reality as sharp as the borders of the more medially located areas. This phenomenon is due to the curvature of the cortex that is highest laterally in the cortex, where auditory areas are located. Flattening artifacts are therefore strongest laterally, where the cortex is curved the most, which consistently results in somewhat fuzzy staining around the lateral most cortical edge. Fuzzy borders of the auditory areas are indeed apparent in flattened sections in Figure 1A. Importantly the fuzziness of the staining still allows for reliable delineation of the auditory areas, and the staining is comparable between wt and ne-Emx2 sections. Further, we compare the quantification of V1 gene markers using un-sectioned whole brains with quantification of V1 derived from 5HT-stained flattened tangential sections (paragraphs six and nine, Results). The quantified sizes of the 5HT-stained V1 were in all cases very similar to the quantifications using marker genes, suggesting that the flattening and staining procure used in this study delivered reproducible results and contained no major artifacts due to the flattening procedure. To emphasize this point more clearly, we have moved the images and graphs of the quantified 5HT-stained flattened sections that were originally figure supplements into Figure 3A and Figure 4A, respectively. Overall we believe that the use of whole-mount and well as sectioned data excludes the possibility that flattening artifacts alter the interpretation of our findings and our conclusions.

We discuss the issue about possible inconsistent delineation of V1 and HO in the comment below.

Another related reservation is the lack of evidence for reliable areal parcellation. Although the experimental design of manipulating the size of V1 is clever and elegant, without reliable parcellation of the cortex, the results fall short of being convincing to demonstrate that the size of V1 is scaled and that there is a clear cause and effect between a change in the size of V1 and the size of neighboring higher order areas.

Apart from the qualitative assessments of V1 and HO in Figures 12, we also report a quantitative assessment of the sizes of V1 and HO in Figure 3 and Figure 4. For this purpose, we use a panel of marker genes that either shows expression domains in V1 alone (Unc5d, Igfbp5) or also in HO (Lmo4, Cad8). All of the reported marker genes have been used successfully in previous studies to assess the size and location of primary areas including V1 (Cholfin and Rubenstein 2007, Chou et al. 2013, Hamasaki et al. 2004, Leingartner et al. 2003, Leingartner et al. 2007, Zembrzycki et al. 2015). Our study is novel in introducing Lmo4 and Cad8 as markers that also can be used to delineate and to quantify HO. We acknowledge that the reviewer’s comment in this regard was appropriate for the original manuscript. We have therefore added a new figure supplement (Figure 3—figure supplement 2) to the manuscript that better illustrates and identifies area borders on the basis of Lmo4 and Cad8 expression in wt and ne-Emx2 brains. In this new figure we show whole brains in dorsal view that were stained in situ for Cad8 or Lmo4. We present two sets of images for wt and ne-Emx2 brains: One showing the raw images and another set showing the in situ images with superimposed annotated area borders on the basis of the marker gene expression domains. We think that these images much better illustrate how area borders were identified and used in this study to quantify V1 and HO sizes in Figure 3 and Figure 4. In addition to the representative cases that are shown on higher magnification in Figure 3 and Figure 4, the new figure supplement provides the reader with an additional set of marker gene-stained whole brains that show comparable area outlines on the basis of marker gene expression domains. We think that incorporating these additional cases provides additional evidence on the reliability and reproducibility of this technique to quantify V1 and HO.

The referees raised several specific issues that were deemed essential. They requested that the revised manuscript demonstrate that: (1) the size of V1 in control and experimental brains was determined in identical fashion;

We emphasize that all specimens were processed identically and the sizes of V1 determined identically on the basis of 5HT staining patterns, area-specific marker gene expression domains, or intrinsic functional response properties, respectively. We again apologize for the inconsistencies in some of the originally presented areal delineations. As we explain in more detail in our response to related comments, we have revised the representative outlines where they were inaccurate.

(2) That the measurements of the borders of VHO are clearly outlined in control and experimental brains;

We apologize for the inconsistencies in the area outlines in the original version of the manuscript. As we have detailed in our response to the specific comments below, we have carefully revised the annotations and have also added a new data as Figure 3—figure supplement 2 that details the gene expression domains of Cad8 and Lmo4 on views of the entire brain and how they relate to area borders in the cortex that we have used for area quantification.

(3) The borders of the extrastriate visual cortex need to be better defined. Does VHO exclude (or include) retrosplenial cortex;

The reviewers’ concern results from the failure of the original manuscript to provide some necessary details on the definition of cortical areas. In Figure 1, we define primary sensory areas and major cortical subdivisions on the basis of 5HT staining on flattened sections of the cortical hemisphere. 5HT is strongly expressed in primary areas V1, S1, auditory areas. A clear change to much lower 5HT staining in the entorhinal cortex makes it possible to identify the border between neocortex and entorhinal cortex laterally. The caudomedial cortical pole is devoid of 5HT staining; hence no distinction between higher order visual areas around V1 and the retrosplenial cortex around the cortical midline can be made on the basis of 5HT staining. Therefore in the qualitative assessment (Figure 1) of the region that lies outside of the primary sensory areas, the retrosplenial cortex is included. To better illustrate cortical parcellation and how they are affected in ne-Emx2 brains, we have added new images showing 5HT staining on the entire cortical hemisphere into Figure 1A. The higher magnification set of images that focuses on extrastriate areas and how they appear to expand qualitatively in ne-Emx2 brains follow after them. We have also revised the text accordingly to now more precisely define the different cortical regions and how we have defined their borders (paragraph one, Results).

In addition, in response to the reviewer’s comment on the borders of HO, and we now present images of the genetic markers Lmo4 and Cad8 and note that the quantitative assessment of V1 and HO sizes were made without including the retrosplenial cortex medially. We have added according statements into the text (paragraph seven, Results) to describe these details more comprehensively.

(4) The lateral border of VHO relative to primary auditory cortex, ventral posterior temporal cortex and rhinal fissure needs to be clearly defined.

We agree that a comprehensive demonstration of how 5HT and marker gene staining relates to these boundaries was missing. To better define these borders using 5HT staining on flattened cortical sections, we have introduced new 5HT images showing the entire cortical hemisphere (Figure 1A). We have also modified the text accordingly to include these definitions (paragraph one, Results).

In addition, to define these borders more clearly in un-sectioned whole-mount brains stained in situ, we have introduced a new figure supplement (Figure 3—figure supplement 1). This figure shows how borders to the auditory cortex and entorhinal cortex were defined through the use of Lmo4 as a marker gene. Due to the curvature of cortex, not all cortical subdivisions are visible from every angle. This is especially true for the auditory cortex and entorhinal cortex in dorsal view. By using three different angles (lateral view, dorsolateral view, dorsal view) of the same Lmo4-stained brain, the new figure supplement now illustrates in raw images and annotated images how sharp gene expression borders were used to define subdivisions of the cortex. For the quantification of HO using marker genes shown in Figure 3 and Figure 4, we have exclusively chosen to use images of the dorsal view, since it is the best suited angle that gives the most-complete view of HO, while still allowing to define the borders to the auditory areas, the entorhinal cortex and the retrosplenial cortex in a single image. We accordingly now refer to this new data in the text (paragraph seven, Results).

This new figure supplement also adds a demonstration of the expression of the gene Lypd1 to the manuscript. We have found that, Lypd1 is not significantly expressed in V1 and HO, but strongly labels the retrosplenial cortex on whole mount images in the dorsal view (Figure 3—figure supplement 1). The gene expression differences of Lmo4 between HO and retrosplenial cortex are more subtle, compared to Lypd1 expression, but detectable (Figure 3—figure supplement 1). By comparing the expression of Lmo4 and Lypd1 in the medial cortex, this figure supplement demonstrates that the subtle Lmo4 gene expression differences in this region reliably reveal the subdivision between HO and the retrosplenial cortex.

More detailed comments directly related to these concerns are elaborated below with recommendations for improvement:

1) The VHO in the EMX2 case includes retrosplenial cortex and extends deeper into temporal cortex than in WT animals. This raises the questions how the borders of VHO were defined and what types of criteria for flattening the cortex were used to compare the sizes of V1 and VHO. There is strong evidence that VHO extends far beyond extrastriate visual cortex. Thus, one option is to drop the "V" and call the region "HO".

We agree with the reviewers’ comment to more conclusively define the borders of V1 and VHO based on 5HT staining in Figure 1. Therefore, we have modified Figure 1A accordingly. We now show 5HT staining also on the entire cortical hemisphere and describe how the borders of 5HT staining in layer 4 reveals major anatomical subdivisions of the cortex including primary sensory area borders and laterally the border from the neocortex to the entorhinal cortex. We have also revised the text accordingly (paragraph one, Results). Further we have followed the reviewers’ suggestion and have dropped the term VHO and instead refer more broadly to higher order areas (HO) throughout the text and figures and now state that the delineation of ‘HO’ in Figure 1 contains higher order visual areas and the retrosplenial cortex, since 5HT staining does not reveal the subdivision between them.

2) Figure 1B. Labeling of the thalamocortical input is challenging. It is difficult to rule out that labeling patterns are produced by injections at particular topographic locations. The experiment should indicate the LGN injection site and the projections in flatmounted cortex, superimposed onto 5HT labeled V1 in the same mouse.

We agree that labeling of TCAs can be challenging, but similar experiments are very common in the field and have been utilized to provide reproducible results. Although it is difficult to rule out entirely that differences in staining patterns in the cortex are influenced by tracer injections into different topographic locations in the thalamus, we believe that the presented DiI stainings provide very solid evidence for the claims made (Figure 1—figure supplement 1): (i) TCAs from the dLG extend more anteriorly in ne-Emx2 brains. (ii) The border between DiI-stained and un-stained cortical tissues remains sharp in ne-Emx2 brains. We believe that these claims are well supported by the presented data, because we observe these staining patterns consistently as demonstrated by examples on five different levels that cover the cortex from medial to lateral. This indicates that the filling of the dLG with dye was broad, robust, and comparable across genotypes. Further, the results of the DiI labeling from the dLG are consistent with an enlarged 5HT-stained V1. We remain confident that expansion of the DiI-stained domains at all tested medial to lateral levels in ne-Emx2 brains are reflecting an enlargement of V1 that is evident using 5HT staining by using an alternative approach.

However, we have made some adjustments to the presentation of this data in the manuscript: We have removed the exemplary single DiI image in Figure 1B and instead in the main text now exclusively refer to the entire set of five different levels that are shown in Figure 1—figure supplement 1 (paragraph one, Results). As suggested by the reviewer, we have also added representative images that show strong and complete filling of the dLG with DiI in the sagittal plane (Figure 1—figure supplement 1, panel B) demonstrating that the filling of the dLG with dye in wt and ne-Emx2 cases is comparable and robust.

Further, we feel that suggested images of DiI-stained, flat-mounted cortical sections with or without 5HT staining would not provide more directly relevant information to support the claims made. After all, we have decided not to introduce such images mainly because the laminar distribution and borders between stained and unstained cortical regions could potentially be distorted due to the flattening procedure. This issue can be circumvented and the laminar DiI distribution clearly demonstrated using sagittal sections as shown in Figure 1—figure supplement 1.

3) Figure 1C. The analysis could be strengthened by 5HT staining of parallel sections showing S1 and V1.

We agree that 5HT staining complementing the apparent shift in distribution of layer 5 output projection neurons in ne-Emx2 sections would be ideal. We would like to emphasize that this experiment requires an extremely long incubation time (4-6-month incubation in PFA at 37°C) in order to achieve sufficient in-vitro retrograde dye-labeling in the cortex from the far out injections sites in the pyramidal decussation and superior colliculus, respectively. In some pilot experiments we indeed generated such dual DiI labelings and performed 5HT stainings after the necessary diffusion time of the tracers using similarly processed specimens. Contrary to clear DiI-staining distributions in the cortex, we never obtained reliable 5HT staining in these brains. Proper immunostainings in these specimens are likely hampered by a combination of many month-long, extreme PFA over-fixation and prolonged elevated storage temperature over a substantial period of time.

Because of the low projected success rate and the fact that the anterior enlargement of 5HT staining in V1 layer 4 in ne-Emx2 brains is very well documented (Hamasaki et al. 2004, Leingartner et al. 2007), we did not intend to perform 5HT staining on consecutive sections for this study. Overall we think that the presented DiI data clearly reveals that in layer 5 in ne-Emx2 brains the balance of corticotectal versus corticospinal projections is altered in a way that is consistent with an anterior expansion of V1 and higher order areas: Qualitatively, ne-Emx2 brains have an enlarged region sending out corticotectal projections, to the expense of a smaller region that send out corticospinal projections.

4) Figure 2A. A more compelling demonstration of the change in the size of V1 and VHO would be to mark reference points in the topographic map, stain the same flattened cortex for 5HT and superimpose the images.

The present Figure 2B is exactly the experiment suggested, in which we marked the borders of V1 in vivo during the creation of the functional maps and then located the marks post mortem on sections stained for 5HT. Fourier imaging reveals the intrinsic functional neuronal properties of neurons in response to a visual stimulus that was moved across the visual field, from left to right and top to bottom and has been shown to reveal the borders of the primary visual area in the mouse (Kalatsky and Stryker 2003).

We should also note that each of the color-coded regions in the response maps represent visual responses to stimulation around a certain region of the visual field. Regions of similar color in different maps can serve as a reference point or reference region for areas responsive to the same portion of the visual field. In the text we now note that the green region representing ~20 to ~30 degrees of elevation/azimuth is clearly enlarged in ne-Emx2 brains, compared to wt brains (paragraph four, Results).

Further, it would strengthen the interpretation of the data if the maps included information about the topographic representations in areas of the lateral extrastriate cortex.

While it might be interesting to demonstrate the complete parcellation of HO into many separate functional areas, doing so would be far beyond the scope of the present manuscript. We chose not to attempt this with the functional maps because we had, and still have, no corresponding genetic parcellation. In addition, the stimuli used for our functional maps like those in Figure 2 were not ideal for demonstrating the organization of all of the extrastriate cortical areas. Hints of them can be seen in at the lower left in the two leftmost panels of Figure 2A, but no attempt was made to delineate them all, as has now been done for many of them in several laboratories (e.g.: Andermann et al. 2011, Garrett et al. 2014).

5) Figure 2B. The cartoon of the mouse thalamus and the labels are inaccurate for the DiI and DiD marked projections. The DiI and DiD injections into WT and ne-Emx2 mice are targeted to different topographic locations of V1 and areas within VHO. These discrepancies weaken the conclusion that the corticothalamic projections are comparable.

We should note that the color-coding of the schematics does not refer to the dye injection sites shown in the original Figure 2B (now Figure 2C). The colors were intended to provide readers that are not familiar with the stereotypic connections between thalamus and cortex with a visual aide using matching colors of the cortical areas and their corresponding thalamic regions to illustrate that S1 is interconnected with the VP, V1 with the dLG, and HO with the PO, respectively. The fact that the reviewers found this color-coding to be confusing led us to remove the color-coding from the schematics in Figure 2. They now are in black and white.

Further, we would like to clarify that it was by design that the injections were made at different topographic locations. By doing so, we aimed to investigate whether the actual connectivity patterns were consistent with the predicted patterns of connectivity based on the 5HT staining around the injection sites. The results indicate just that: Following overall changes in V1 size in ne-Emx2 brains, the area borders of V1 and HO shift anteriorly. This demonstrates that although parts of V1 and HO in ne-Emx2 brains are located at different topographic locations on the cortical sheet compared to wt, the neurons show connectivity patterns that are consistent with their altered intrinsic areal identity, not their topographic location on the cortical sheet. We have revised the text section to emphasize this issue more clearly (paragraph five, Results).

6) Figure 3B. The borders of VHO in WT and ne-Emx2 mice are different. Again VHO is ill-defined and its borders are not consistently outlined. For example, the Cad8 (WT) stained region around the tip and continuing along the medial border of V1 is much larger than indicated by the stippled line.

We thank the reviewers for bringing this important point to our attention. We did not outline the gene expression domains with enough accuracy and consistency in the figures of the original manuscript, and we apologize for that. We have carefully revised the outlines that in the current manuscript version much more accurately delineate the gene expression borders. Moreover we hope that the more detailed definition of HO mentioned above (detailed comments #1) together with the newly added whole mount gene expression images (Figure 3—figure supplement 12) that show additional Cad8- and Lmo4-stained whole brain cases leads to a more conclusive definition of the gene expression borders and how V1 and HO sizes were determined. As mentioned above, we have modified the text accordingly to emphasize this issue (paragraph seven, Results).

In addition, V1 in ne-Emx2 mice has an unusual shape with an unexpected "tipped nose", indicating that the region enclosed by the stippled line may be larger than V1. The flattening technique can contribute to an overestimation of the size differences reported in the study.

The comment of the reviewers is correct. In ne-Emx2 brains the shape of V1 is slightly affected. This is mostly apparent around the anterior tip of V1. The revised outlines now more accurately match the expression domains of the probed genetic markers. We have clarified in the revised manuscript that the measurements in Figure 3 and Figure 4 were made on un-sectioned whole brains and not on flattened cortical sections. Although, performing in situ hybridizations using whole brains is more difficult, compared to in situ hybridizations on sections, we have used the whole brain approach for quantification purposes to prevent potential sizing artifacts due to the flattening and cutting procedures that were mentioned by the reviewers. The images in Figure 3 and Figure 4 show the staining patterns on the intact brain and we therefore remain confident that the quantifications are accurate and reproducible. In the revised manuscript, we more clearly state that whole un-sectioned whole brains were used for the quantifications to prevent confusion of the reader regarding this point (paragraph six, Results).

7) Figure 4B, similar to Figure 3B. Cad8 (WT) and Lmo4 (WT) include weakly stained regions in medial extrastriate cortex, which are not included in cKO mice.

As mentioned in more detail in our reply to the reviewers’ comment #6 above, we again appreciate the reviewers’ attention to important detail and for bringing this important issue to our attention. We apologize for the inaccurate outlines and have revised them carefully.

[Editors' note: further revisions were requested prior to acceptance, as described below.]

Reviewer #1:

[…] One remaining point: The Abstract needs to be slightly rewritten to bring it into line with the rest of the paper. In the Abstract it is stated: "we have generated bi-directional changes in primary visual cortex size in vivo and have used it as a model to show a novel and prominent function for genetically determined primary visual area size, which also proportionally dictates the sizes of higher order visual areas." It seems (again) that the authors want to say that V1 size controls the size of surrounding higher order areas. Previous review suggested there was not enough evidence for such a conclusion, and pointed to an alternate interpretation: genetic factors intrinsic to cortex control the size of both V1 and HO areas in coordination. The cover letter for the revised MS and the new Discussion section appear to agree with the latter conclusion.

We agree to the statement of the reviewer. Our data supports the claim that genetic factors intrinsic to the cortex control the size of both V1 and HO areas in coordination. We have modified the Abstract accordingly to bring it in line with the rest of the manuscript.

Reviewer #3:

Although V1 and the organization of surrounding cortex are beautifully illustrated in several newly added images, there are remaining concerns that the revision failed to analyze the outstanding material in a consistent fashion. This impression is based on the inexplicable discrepancy of the definition of HO shown in Figure 1A and Figure 3A, which indicates that the authors are unsure how to subdivide the real estate. In Figure 1A it appears that the border of HO runs from the anterior tip of retrosplenial cortex along the posterior border of S1, continues along the border of auditory cortex and ends by intersecting the rhinal fissure. This does not fit the scheme shown in Figure 3A, in which HO does not extend to the border of auditory cortex and does not include retrosplenial cortex. If I understand this correctly, HO in Figure 3A is clearly smaller than in Figure 1A and does not include ventral posterior temporal cortex nor does it include retrosplenial cortex. In fact, HO Figure 3A is what was measured in Figure 3 and Figure 4. Maybe these discrepancies were simply an oversight which can be easily corrected. But the lapse heightened my concerns that the size of HO was inconsistently assessed (e.g. in Figure 3B the border of Cdh8-expression in wt is much more medial than indicated and differs from the border in ne-Emx2) and that the substance of the paper is less solid than suggested by the graphs shown in Figure 3B and Figure 4B. I recommend re-analyzing the material with a more critical eye on the detailed borders of HO. I am sure that the authors are acutely aware that details matter a lot more in small than large brains.

We would like to mention that we have analyzed the data in a consistent fashion. In general, claims that we have made about the HO as defined in Figure 1 are qualitative (e.g. HO appear enlarged in ne-Emx2 brains), whereas the measurements of HO in Figure 3 were made without including the retrosplenial cortex.

The reviewer’s concern reflects that our revised manuscript failed to better explain how HO areas reflect slightly different cortical territories in Figure 1, compared to Figure 3 and why this is the case. We have revised the manuscript to better state this discrepancy and why and have added some new data that better explains why we exclude retrosplenial cortex from our quantifications in Figure 3 and Figure 4 (details are explained below).

In Figure 1 we have used 5HT staining to introduce the central issue of the paper: Upon genetically increasing V1 size, the surrounding higher order areas appear to be qualitatively enlarged as well. We have chosen to use 5HT staining to introduce this central issue in order to present the analysis in a way that is comparable to virtually all studies that have analyzed and demonstrated primary area patterning by using 5HT staining as a ‘gold standard’ of the cortical areal layout and to extend on them in a consistent fashion by highlighting altered higher order areas by using the same technique. Our main focus is using V1 as a model to probe the influence of altered V1 size on the proportions of higher order visual areas. An issue that is not understood. As mentioned in our initial review comments, the caveat is that 5HT staining does not reveal the retrosplenial cortex and hence the HO complex that we needed to define initially in Figure 1 to set the stage for the rest of the data includes it. This important consideration is clearly stated to the potential readership of the manuscript in the definition that can be found in paragraph one, Results.

The dilemma, which might lie on the bottom of some of the reviewer’s concern, might be that especially in the mouse, there is no commonly agreed upon or long-established terminology for the cortical region of interest. At least this is the case for the rodent brain. Descriptions in published studies are in itself inconsistent ranging from ‘extrastriate cortex’ to ‘higher order visual areas’ and ‘higher order cortex’. All of these previously used terms are slightly inaccurate. For example, there is no such thing as a striate cortex in rodents, but it is common to describe higher order visual areas in rodents as such since there appears to be a general agreement that extrastriate areas in rodents reflect higher order areas around V1 and that these areas are analogous to the striated/extrastriated areas that are characteristic for higher, more complex mammalian brains like cats, monkeys, and humans.

In the initial manuscript version we therefore referred to this cortical territory as higher order visual areas (VHO), which was felt to be incorrect. We agreed upon this view and that it would be more appropriate to describe this territory more broadly as higher order area complex (HO) throughout and define better which cortical subdivisions are included and excluded in Figure 1 versus Figure 3 and Figure 4.

Compared to 5HT staining in Figure 1, we in Figure 3 use genetic markers that enabled us to distinguish between higher order visual areas medially of V1 and the retrosplenial cortex around the cortical midline. In this quantitative assessment, we have therefore excluded the retrosplenial cortex from our definition of HO since genetic markers enabled us to make more precise anatomical claims about the core areas of interest (mainly higher order visual areas) that were assessed quantitatively in Figure 3 and Figure 4. The main focus of our study is genetic scaling between primary visual cortex and higher order visual areas. Also, due the fact that the retrosplenial cortex is not a higher order cortical area (Garrett et al. 2014, Kalatsky and Stryker 2003, Marshel et al. 2011, Vann, Aggleton, and Maguire 2009) and due to unpublished data from our lab (see below for details), we had reasons to exclude the retrosplenial cortex from the quantifications of HO in this study. Overall we do believe that the claims of the manuscript are solid and valid, although HO in Figure 3 and Figure 4 reflect a smaller cortical territory, compared to Figure 1, because the anatomical cortical subdivisions are accurately described in each case. We have nonetheless made the attempt to improve the clarity of the text and have revised the manuscript to more explicitly state that the definitions of HO in Figure 1 and Figure 3 and Figure 4 are slightly different (paragraphs one, seven and eight, Results).

We also have decided to include an additional piece of unpublished data into the manuscript that elaborates specifically on the scaling of the retrosplenial cortex and how it differs from higher order visual areas as measured by Lmo4 and Cdh8 staining: To complement the Lypd1 WMISH staining in wt brains, we have added Lypd1 WMISH staining on ne-Emx2 brains into Figure 3—figure supplement 1 and have quantified it. The Lypd1 gene expression domain around the territory of the retrosplenial cortex is significantly larger in ne-Emx2 brains (114.3% ± 5.2%, p = 0.0225, n = 4), compared to wt brains. But importantly, this size increase is NOT proportionate to the size increases of V1 (~145%) and HO as measured by Lmo4 and Cdh8 (~145%) staining reported in Figure 3. This suggests that increased V1 size is specifically accompanied by a proportionate size increase of related higher order visual areas. This new data explains why we have avoided including the retrosplenial cortex area into the quantitative assessment of higher order visual areas. This new data also helps to more precisely define the central finding of the manuscript: In the revised manuscript we now state that genetic mechanisms control the linear scaling between RELATED cortical primary and higher order sensory areas (e.g. vision). To incorporate these changes and to bring the manuscript in line with the presented data, we have modified the title slightly, have added a passage describing the new data (paragraph seven, Results) and refer to Cdh8/Lmo4 labeled HO as ‘related HO’ or ‘higher order visual areas’ throughout the main text and Discussion.

The schematic in Figure 3 was intended to serve as a visual aide to show location and approximate perimeters of primary and higher order visual areas as defined previously (Wang and Burkhalter 2007) and their overall similarity to the expression domains of Cdh8 and Lmo4 surrounding V1. We have removed the schematic from the revised Figure 3A.

[Editors' note: further revisions were requested prior to acceptance, as described below.]

Reviewer #3:

1) To resolve the conflict between the measured HO (Figure 3, Figure 4) and HO defined by 5HT staining (or the virtual absence of 5HT expression) in Figure 1, I suggest renaming it HO-5HT to indicate the conditions under which the observation was made and unequivocally state that HO and HO-5HT are not the same.

We thank the reviewer for the constructive comment and suggestion to resolve potential misunderstandings about slightly different definitions of higher order areas derived from stainings using different sets of markers. We have followed the suggestion and have termed the HO as stained by 5HT as HO-5HT throughout the text, in Figure 1 and its figure legend. Consistently, we have also amended the annotation of HO as labeled by Rorb in Figure 1 as HO-Rorb in Figure 1 and its legend.

2) HO includes at a minimum visually dominated as well as higher order auditory areas. I therefore recommend revising the sentence (Abstract) to: "… surrounding higher order sensory areas". I further recommend that similar revisions be made throughout the text to make it absolutely clear that the cortex measured here is more extensive than extrastriate visual cortex.

We thank the reviewer for the recommendation. We would like to clarify that the complex of higher order areas that was quantified using Lmo4 and Cdh8 marker genes does not include primary auditory cortex or secondary auditory areas. As it is clearly shown in Figure 3—figure supplement 1 A and B (as an example using Lmo4 staining), primary and secondary auditory areas are included in the complex of auditory areas that we have termed “Aud”. As it is evident in three different angles in Figure 3—figure supplement 1, this “Aud” complex is excluded from our measurements. We have currently another paper in review that in high detail is concerned about the scaling of the auditory areas in comprehensive sets of different inbred mouse strains and transcription factor mutants. Unfortunately it is too premature to reference at this point, but we would like to emphasize that we have solid evidence about how primary and secondary auditory areas relate to expression sub-domains of Lmo4 and other area markers that are expressed in this cortical region in addition to the well documented and clearly stated exclusion of secondary auditory areas (defined as ‘Aud’, e.g. Figure 3—figure supplement 1) that are already present in the manuscript. Considering these points we would like to re-emphasize that secondary auditory areas are excluded from the presented measurements and that we remain confident that our claims (e.g. Abstract) are in line with the presented data.

3) I recommend revising the border of HO (wt/Cdh8) in Figure 3B. It is clearly evident in the image that the Cdh8 expression includes a wedge medial to V1 which extends to the tip of the arrow head. A similar adjustment of the medial border is necessary for ne-Emx2 Cdh8 (Figure 3B). The same revisions are necessary in Figure 4B in which the medial borders of Cdh8 expression in wt and cKO are more medial than indicated.

We would like to acknowledge that the reviewer has an exceptional eye and commitment to detail. We agree that around the area in question (around arrowheads in Figure 3), there is some Cdh8 staining detectable. However, we would like to mention that this gene expression domain shows much lower staining compared to the outlined complex of all the other regions around V1 that we have quantified (please re-examine Cdh8 whole brain images in Figure 3—figure supplement 2B). We have mentioned this issue consistently in all previous versions of the manuscript as a discernable difference between the expression domains of Lmo4 and Cdh8 (arrows in Figure 3, see paragraph seven, Results): Due to much lower expression compared to all the other more lateral regions around V1 (e.g. Figure 3—figure supplement 2B), we have excluded this domain from the measurements and consistently from the outlines. Due to these criteria we think that the present outlines are well defined on the basis of similarly high Cdh8 gene expression levels, compared to detectable but much lower levels in the region of question. We feel that modifying the outlines to include this area of lower Cdh8 expression could potentially appear inconsistent to potential readers and would also require us to comment on this issue and criteria (that compared to as is, would not be based on similarly high Cdh8 expression levels) in much more detail in the text. Since this issue has always been well documented (arrowheads in Figure 3) and mentioned in the text (paragraph seven, Results) by comparing Lmo4 and Cdh8 expression levels in this region, we have the impression that altering the outlines would not be beneficial. To the contrary, it would require us to re-define gene expressions areas without adding additional insights to the conclusions of the manuscript. We have revised the text slightly for clarity to emphasize more progressively that the excluded region does in fact expresses Cdh8, but at much lower levels, compare to the outlined and defined area complex of consistently much higher Cdh8 gene expression (paragraph seven, Results).

https://doi.org/10.7554/eLife.11416.015

Article and author information

Author details

  1. Andreas Zembrzycki

    1. Molecular Neurobiology Laboratory, The Salk Institute for Biological Studies, La Jolla, United States
    Contribution
    AZ, Conception and design, Acquisition of data, Analysis and interpretation of data, Drafting or revising the article
    Contributed equally with
    Adam M Stocker
    For correspondence
    1. azembrzycki@salk.edu
    Competing interests
    The authors declare that no competing interests exist.
    ORCID icon 0000-0001-8468-4195
  2. Adam M Stocker

    1. Molecular Neurobiology Laboratory, The Salk Institute for Biological Studies, La Jolla, United States
    Present address
    1. Biosciences Department, Minnesota State University, Moorhead, United States
    Contribution
    AMS, Acquisition of data, Analysis and interpretation of data, Drafting or revising the article
    Contributed equally with
    Andreas Zembrzycki
    Competing interests
    The authors declare that no competing interests exist.
  3. Axel Leingärtner

    1. Molecular Neurobiology Laboratory, The Salk Institute for Biological Studies, La Jolla, United States
    Present address
    1. University Cancer Center Hamburg, University Medical Center, Hamburg, Germany
    Contribution
    AL, Acquisition of data, Analysis and interpretation of data
    Competing interests
    The authors declare that no competing interests exist.
  4. Setsuko Sahara

    1. Molecular Neurobiology Laboratory, The Salk Institute for Biological Studies, La Jolla, United States
    Present address
    1. MRC Centre for Developmental Neurobiology, Kings College, London, United Kingdom
    Contribution
    SS, Acquisition of data, Analysis and interpretation of data
    Competing interests
    The authors declare that no competing interests exist.
  5. Shen-Ju Chou

    1. Molecular Neurobiology Laboratory, The Salk Institute for Biological Studies, La Jolla, United States
    Present address
    1. Institute of Cellular and Organismic Biology, Academia Sinica, Taipei, Taiwan
    Contribution
    SJC, Acquisition of data, Analysis and interpretation of data
    Competing interests
    The authors declare that no competing interests exist.
  6. Valery Kalatsky

    1. Center for Integrative Neuroscience, Department of Physiology, University of California, San Francsisco, San Francisco, United States
    Present address
    1. Enthought Inc, Austin, United States
    Contribution
    VK, Acquisition of data, Analysis and interpretation of data
    Competing interests
    The authors declare that no competing interests exist.
  7. Scott R May

    1. Molecular Neurobiology Laboratory, The Salk Institute for Biological Studies, La Jolla, United States
    Contribution
    SRM, Acquisition of data, Analysis and interpretation of data
    Competing interests
    The authors declare that no competing interests exist.
  8. Michael P Stryker

    1. Center for Integrative Neuroscience, Department of Physiology, University of California, San Francsisco, San Francisco, United States
    Contribution
    MPS, Analysis and interpretation of data, Drafting or revising the article
    Competing interests
    The authors declare that no competing interests exist.
  9. Dennis DM O'Leary

    1. Molecular Neurobiology Laboratory, The Salk Institute for Biological Studies, La Jolla, United States
    Contribution
    DDMO'L, Conception and design, Analysis and interpretation of data, Drafting or revising the article
    Competing interests
    The authors declare that no competing interests exist.

Funding

National Institutes of Health (NS31558)

  • Dennis DM O'Leary

National Institutes of Health (MH086147)

  • Dennis DM O'Leary

National Institutes of Health (EY02874)

  • Michael P Stryker

The funders had no role in study design, data collection and interpretation, or the decision to submit the work for publication.

Acknowledgements

We thank Kevin Jones for providing Emx1-IRES-CRE mice. This work was supported by the Vincent J Coates Chair of Molecular Neurobiology at the Salk Institute for Biological Sciences (DDMO).

Ethics

Animal experimentation: All experiments were approved under Protocol #09-012 and conducted following the guidelines of the Institutional Animal Care and Use Committee at the Salk Institute and were in full compliance with the guidelines of the National Institutes of Health for the care and use of laboratory animals.

Reviewing Editor

  1. Moses V Chao, Reviewing Editor, New York University School of Medicine, United States

Publication history

  1. Received: September 5, 2015
  2. Accepted: December 23, 2015
  3. Accepted Manuscript published: December 24, 2015 (version 1)
  4. Version of Record published: January 26, 2016 (version 2)

Copyright

© 2015, Zembrzycki et al.

This article is distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use and redistribution provided that the original author and source are credited.

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