Reticulon 4a/NogoA locates to regions of high membrane curvature and may have a role in nuclear envelope growth

Reticulon 4a (Rtn4a) is a membrane protein that shapes tubules of the endoplasmic reticulum (ER). The ER is attached to the nuclear envelope (NE) during interphase and has a role in post mitotic/meiotic NE reassembly. We speculated that Rtn4a has a role in NE dynamics. Using immuno-electron microscopy we found that Rtn4a is located at junctions between membranes in the cytoplasm, and between cytoplasmic membranes and the outer nuclear membrane in growing Xenopus oocyte nuclei. We found that during NE assembly in Xenopus egg extracts, Rtn4a localises to the edges of membranes that are flattening onto the chromatin. These results demonstrate that Rtn4a locates to regions of high membrane curvature in the ER and the assembling NE. Previously it was shown that incubation of egg extracts with antibodies against Rtn4a caused ER to form into large vesicles instead of tubules. To test whether Rtn4a contributes to NE assembly, we added the same Rtn4a antibody to nuclear assembly reactions. Chromatin was enclosed by membranes containing nuclear pore complexes, but nuclei did not grow. Instead large sacs of ER membranes attached to, but did not integrate into the NE. It is possible therefore that Rtn4a may have a role in NE assembly.
1
Introduction
The nuclear envelope (NE) encloses the nucleus (Hetzer et al., 2005; Goldberg, 2004) and consists of two parallel sheets of membrane connected at nuclear pore complexes (NPCs). The outer membrane is continuous with the rough endoplasmic reticulum (ER). During mitosis and meiosis in higher eukaryotes the NE is dismantled. The lamina and NPCs are solubilised and the membranes disperse into the cytoplasm by retraction into the ER (Ellenberg et al., 1997; Yang et al., 1997) or by vesiculation (Vigers and Lohka, 1991).
During telophase the NE is reassembled around chromosomes in a multistage process, involving accumulation of different membrane populations and NE and NPC proteins. Evidence from Xenopus egg extracts suggest that there are least two vesicle populations (Vigers and Lohka, 1991; Macaulay and Forbes, 1996; Drummond et al., 1999) which fuse to form tubules and sheets (Wiese et al., 1997; Goldberg et al., 1992; Hetzer et al., 2001) which enclose the chromatin. Fusion of nuclear membranes requires hydrolysis of GTP (Macaulay and Forbes, 1996) by Ran (Hetzer et al., 2000; Zhang and Clarke, 2000; Zhang et al., 2002) and also the p97–UFD1–NPL4 complex (Hetzer et al., 2001). The mechanisms of targeting and fusion are unknown. NPCs are apparent after a few minutes in egg extracts (Goldberg et al., 1992), but NPC proteins accumulate in a temporal order in culture cells (Bodoor et al., 1999) as do structural intermediates in the NPC assembly process (Goldberg et al., 1997; Kiseleva et al., 2001).
During S-phase the NE grows as the DNA content in the nucleus is increased (Winey et al., 1997). The NE also grows during oogenesis. Although interphase NE growth is often considered separately from telophase, there are common features. Conceptually, once the chromatin is enclosed in telophase, there is no obvious difference between the subsequent growth phase and interphase growth. Like enclosure, growth requires the AAA-ATPase p97, but instead of UFD1 and NPL4 it is complexed with p47 (Hetzer et al., 2001), suggesting the mechanisms are related but distinct.
Rtn4a/NogoA (hereafter referred to as Rtn4a) is a member of reticulon family and is one of three splice variants of the RTN4 gene (Oertle and Schwab, 2003). It has attracted much interest recently because of its inhibitory role in neurite outgrowth (Prinjha et al., 2000; Yan et al., 2006). Rtn4a has also been shown to localise to the ER (van de Velde et al., 1994) but is restricted to the tubular network and excluded from the peripheral ER sheets and NE (Voeltz et al., 2006). Rtn4a was shown to be required for formation of ER tubules from sheets and vesicles (Voeltz et al., 2006) and it was suggested that reticulons could induce and stabilise the high curvature of the membrane required to maintain tubules. It is possible that the unusual topography of the reticulons, with their long putative transmembrane domains, could induce curvature when clustered.
Rtn4a does not appear to be concentrated at the NE during interphase (Voeltz et al., 2006) possibly because the NE membrane bilayers consist of flat sheets with low membrane curvature. However, nuclear assembly involves considerable reorganisation of membrane topology. It is thought that ER tubules, shaped by Rtn4a (Voeltz et al., 2006), feed into the NE (Ellenberg et al., 1997) and vesicles may also contribute (Vigers and Lohka, 1991; Liu et al., 2003; Prunuske et al., 2005). These vesicles and tubules have to be converted to a large flattened double sheet during NE assembly. Recently, it was shown that Rtn4a may have an essential role in NE disassembly in C. elegans (Audhya et al., 2007). Therefore, we decided to test if Rtn4a could have a role in NE formation, both during interphase and telophase. We used a high resolution surface imaging technique, field emission in-lens scanning electron microscopy (feiSEM1) to look at the structure of interphase growing NEs during Xenopus oogenesis and in telophase in vitro. We show that highly curved membrane regions of the forming or growing NEs preferentially contain Rtn4a. Such regions include the junctions between apparently fusing vesicles and the edges of flattening membranes. An antibody against Rtn4a was also shown to affect NE assembly. We therefore suggest that Rtn4a could have a role in NE formation and growth.
2
Results
The ER connects to the NE in interphase and has been implicated in the post mitotic assembly of the NE (Ellenberg et al., 1997; Yang et al., 1997; Mattaj, 2004). Much of the interphase ER forms a tubular network which appears to feed into the NE during reassembly (Ellenberg et al., 1997; Hetzer et al., 2001). Rtn4a has been found to be involved in shaping the ER into tubules in specific regions (Voeltz et al., 2006). Therefore, we speculated that if the ER needs to be in a tubular form to contribute to NE growth and assembly then Rtn4a may have a role. First we asked whether Rtn4a is associated with membranes that are contributing to NE growth, and then we investigated whether Rtn4a might be required for NE assembly.
2.1
Rtn4a is located at inter-membrane junctions between cytoplasmic vesicles near the NE
During oogenesis in Xenopus, oocytes are arrested in pre-prophase when the nucleus grows to over 100 μM diameter. The NE must expand by the addition of membranes and assembly of lamina and NPCs. We previously observed, both in thin section TEM of whole oocytes and isolated nuclei and in feiSEM of isolated NEs, that growing stage III oocytes have more extraneous ER-like membranes associated with the NE than mature stage VI (Morozova and Kiseleva, 2006).
FeiSEM analysis of the extraneous membranes localised near the NE showed that many were present as structures that look like long lines of inter-connected vesicles (Fig. 1a). We are not certain of the origin of these structures but their surface has an ER-like appearance, with ribosome-like particles on the surface (Fig. 1b and d, arrowheads). The vesicle-like structures are joined together by a short thin tubular connection of ∼20 nm diameter (Fig. 1b, white arrows). The same structures were also observed in TEM thin sections of whole oocytes, showing that they are not artifacts of NE isolation or feiSEM specimen preparation (Fig. 1c). We have also used different fixation methods (see Section 4.1). In TEM sections (Fig. 1c) the 20 nm diameter inter-connecting tubes are continuous with the vesicle membranes and therefore appear to be membrane bridges between vesicle-like structures.
Anti-Rtn4a immuno-gold labelling with a previously characterised affinity purified anti Xenopus Rtn4a antibody (Voeltz et al., 2006) showed that Rtn4a was present on the surface of vesicles (Fig. 1f). In the inter-connected vesicle structures, Rtn4a labelling was concentrated at the junctions between connected membrane structures where the membrane bridge is located (Fig. 1d and e). Inter-connected larger membrane structures were also observed (Fig. 1g and h) where Rtn4a was located at the junction between them. Immuno-gold labelling of TEM sections of whole oocytes also shows localisation of Rtn4a at the contact point of adjacent vesicles. (Fig. 1i). Rtn4a therefore appears to locate to the inter-connections between membrane structures (Fig. 1j). These results show that Rtn4a locates to specific regions on NE associated cytoplasmic membranes.
2.2
Rtn4a is present on membranes attached to growing NEs
We isolated NEs from stage III oocytes and immuno-gold labelled them for Rtn4a. There was some labelling of the ONM (Fig. 2a) but membrane structures associated with the NE were heavily labelled and in some cases, in a specific pattern. Fig. 2a is an image of the surface of a stage III NE showing NPCs, rough (ribosome-containing) vesicles (RV), smooth (ribosome-free) vesicles (SV), and rough membranes that appear to be flattening onto the ONM (FM). We see that there is labelling of the ONM and vesicles are labelled to a varying degree. Smooth vesicles are not labelled, whereas rough ER type vesicles are. This shows that Rtn4a is associated with some but not all membranes that are associated with the ONM of a growing NE.
2.3
Rtn4a localises to the junction between the ONM and membranes attached to it
Membrane structures could be seen attached to the ONM. Rtn4a appeared to be located near the point of contact between the membrane structures and the ONM (Fig. 2b–d). Therefore, it appears that Rtn4a accumulates both at junctions between ER-like membranes and between NE associated membranes that are attached to the ONM. NEs labelled with an antibody (CEL5C) to the ER protein ribophorin (Drummond et al., 1999) showed a more even distribution over flattened membranes, some vesicles and the outer nuclear membrane (Fig. 2e).
2.4
Rtn4a localises to the edges of flattened membranes
Some of the observed NE associated membranes had the appearance of collapsed spheres (Fig. 3a) whereas many are flattened (Fig. 3b–f). It is not always clear from feiSEM images whether such membranes are fused to the ONM or simply lying on top, so thin section TEM was carried out and showed continuity between the ONM and overlying flattened membrane structures (Fig. 4, black arrow). Points of contact between the ONM and cytoplasmic membrane structures were also observed (Fig. 4, white arrows). Potentially corresponding images were seen by feiSEM, in which the flattened membrane structures appear continuous with the NPC-containing ONM (Fig. 3b, arrows).
Using immuno-gold labelling, we observed that Rtn4a appeared to locate preferentially around the edges of the flattened membranes (Fig. 3d–f). This was quantified by counting the number of gold labels that were within 30 nm of the edge compared to greater than 30 nm from the edge (Fig. 5), which gave a ratio of approximately 2:1, for edge compared to the interior. This was in contrast to the ER protein, ribophorin, which was distributed more away from the edge (Fig. 5). The preferred edge location suggests that Rtn4a tends to locate or accumulate at regions with the highest curvature. We conclude that Rtn4a marks the highly curved edges of flattened sheets of presumed ER membrane attached to the ONM.
2.5
Rtn4a is present at the edges of membranes that are flattening onto the chromatin
During telophase, membranes also flatten onto chromatin (Goldberg et al., 1992; Macaulay and Forbes, 1996). We wanted to test if the edge location of Rtn4a that we observed in growing oocyte NEs also occurred in chromatin bound flattening membranes. To do this, we used a cell free system from Xenopus eggs which can be used to assemble nuclei in vitro (Goldberg and Allen, 1993; Lohka and Masui, 1984). Nuclei were isolated from assembly reactions, fixed and immuno-gold labelled for Rtn4a. At early stages (2–5 min) vesicles bind to the chromatin and flatten (Goldberg et al., 1992; Wiese et al., 1997). In such vesicles, Rtn4a localises preferentially around the edges (Fig. 6a and b), compared to a general ER protein, ribophorin (antibody CEL5C—Drummond et al., 1999), which was more randomly distributed (Fig. 6d). To show this formally we counted the distribution of gold particles located within 30 nm of the vesicle edge, or further away in 30 nm increments, using images from three separate experiments (Fig. 6e).This shows that Rtn4a, compared to ribophorin, is preferentially located near the membrane edge. Therefore, as in growing oocytes, Rtn4a preferentially locates to the edges of flattening membranes. At later stages of assembly when the chromatin is enclosed, and there are no longer any highly curved membrane edges (except at the nuclear pores), the Rtn4a distribution on the NE appeared random and low level (Fig. 6c). This is consistent with Rtn4a’s preference for curved membranes (Voeltz et al., 2006). Although there are highly curved membranes in the nuclear pores we see no Rtn4a labelling there. The high membrane curvature at the NPC might be maintained by nucleoporins which could exclude the accumulation of Rtn4a, or it is possible that Rtn4a is present but not detected by the antibody due to epitope masking.
2.6
Disrupting Rtn4a prevents NE growth
Although Rtn4a is not preferentially located to the NE during interphase (Voeltz et al., 2006), our results in stage III oocytes show that it does locate to membranes that are attached to the interphase NE and suggested the possibility that it may be involved in NE assembly or growth. Therefore, we wanted to investigate if Rtn4a might be required for NE assembly. We incubated egg extracts with a previously characterised anti Xenopus Rtn4a antibody directed against the cytoplasmic facing N-terminus (Voeltz et al., 2006) which is specific to Rtn4a and not present in other Rtn4 spliced variants or other reticulon proteins.
This antibody inhibits the formation of ER tubules in similar egg extracts (Voeltz et al., 2006) and likewise we found that it inhibited ER tubule formation. Instead, ER membranes assembled into large vesicular structures (Fig. 7b) rather than tubules as seen in the no-antibody control reactions (Fig. 7a). These large membrane structures are ER-derived because they have ribosomes on their surface. This shows that the anti-Rtn4a antibody had a dominant effect on the formation of ER tubules.
To determine if the Rtn4a antibody affected NE assembly, the same egg extracts were incubated with the antibody (see Section 4.1) or with buffer or with an irrelevant antibody, before adding sperm chromatin to initiate nuclear assembly. Controls showed rapid binding, fusion and flattening of vesicles onto chromatin, following by chromatin decondensation, enclosure and NE growth to form large roughly spherical nuclei >10 microns in diameter, as expected (Goldberg et al., 1997; Wiese et al., 1997 and Fig. 7c). In the presence of the Rtn4a antibody we found that nuclei did assemble with an apparently completely enclosed unbroken NE (Fig. 7d–g) that had apparently normal NPCs (Fig. 7i, black arrows). However, the chromatin failed to decondense and remained as a dense sperm-shaped object surrounded by NE (Fig. 7d–g). To quantify this apparent NE growth defect, we measured the two-dimensional area occupied by each nucleus (excluding attached membranes, see below) as viewed from above by feiSEM (Fig. 7h). Nuclei in Rtn4a antibody-inhibited reactions had ∼80% reduced area compared to controls, confirming the growth defect.
There were in addition, large (several microns) membrane structures extending from the nucleus (Fig. 7d–g, arrows). The extensions are clearly rough ER-like and contain ribosomes (Fig. 7i, arrowheads). There is a sharp demarcation between the chromatin attached membrane which contains NPCs and the membrane extensions which contain ribosomes but not NPCs (Fig. 7i, white arrows). The extensions are clearly continuous with the NPC-containing NE. Therefore, it appears that perturbing Rtn4a does not prevent enclosure of the chromatin by NE. However, the chromatin and NE do not expand despite the attachment of large ER-like sacs to the ONM. These sacs are similar to the large ER vesicle formed in the cytosol in the presence of the antibody, except they are attached to the NE.
Antibodies and reagents against other NE and ER proteins do not have such an affect on nuclear assembly or ER tubulation. Addition of wheat germ agglutinin, which binds to certain nucleoporins (Hanover et al., 1987), to extracts results in nuclei without NPCs but it has no effect on ER tubulation and does not result in membrane extensions (Goldberg et al., 1997). Addition of anti-nucleoporin antibodies (Mab414 and QE5) (unpublished results) or depletion of extracts with antibodies to specific nucleoporins, Nup214 (Walther et al., 2002) or Nup153 (Walther et al., 2001), also affected NPC structure but again does not affect the ER or NE membranes. Depletion of lamin B3 resulted in small spherical nuclei with normal NPCs but had no obvious effect on ER tubules or membrane extensions (Goldberg et al., 1995). The CEL5C antibody against the ER protein ribophorin (Drummond et al., 1999) also did not affect tubulation or membrane extensions when added to extracts (unpublished result). Similar experiments were also done (Voeltz et al., 2006) using antibodies to IP3R and TRAPα which also did not affect ER tubule formation. Therefore, we believe that the growth defects and extensions are a specific effect of the anti-Rtn4a antibody used in this study.
3
Discussion
We have found that Rtn4a is localised to junctions between membrane structures and at the edges of flattened membranes associated with growing NEs both in oocytes and in nuclei assembling in vitro. This is the first direct evidence to support the proposal that Rtn4a locates to regions of high curvature (Voeltz et al., 2006). It is a unique localisation that has not been seen for other NE or ER proteins. We believe this localisation may be driven by Rtn4a itself rather than by interacting with other proteins because high level over-expression of Rtn4a in COS cells (Voeltz et al., 2006) and the related Rtn1 in yeast (De Craene et al., 2006) does not change the proteins’ localisations and therefore is unlikely to rely on other titratable factors. The location of Rtn4a at the junction between cytoplasmic membranes and NE membranes suggested to us the possibility that Rtn4a may also have a role in NE assembly.
Addition of the Rtn4a antibody to nuclear assembly reactions allowed NE formation in egg extracts but chromatin remained condensed and the NE did not expand. Therefore, the antibody has a dominant effect on the growth phase of NE assembly but not on the initial enclosure and NPC assembly. As Rtn4a is an integral membrane protein that appears to be involved in shaping membranes (Voeltz et al., 2006) we suggest that the effect of the antibody is to perturb nuclear membrane dynamics and assembly, as shown previously for the ER (Voeltz et al., 2006).
One possible speculation for the localisation of Rtn4a at membrane–membrane junctions in oocytes is that it may take part in fusion or stabilisation of membrane curvature during fusion. Inter-membrane fusion involves transitory extremes of curvature involving membrane stalks between vesicles (Smeijers et al., 2006; Yang and Huang, 2002). Rtn4a could be involved in stalk formation or stabilisation in certain NE membranes. We do not know if these membranes in oocytes are actively fusing membranes or if they are more stable or intermediate structures. Rtn4a could be involved in stabilising the conformation of these junctions by maintaining the high membrane curvature to facilitate membrane flow into the NE.
Our in vitro antibody inhibition experiments suggest that ER-like membranes can attach to the NE when Rtn4a is perturbed but the NE fails to grow. When Rtn4a is perturbed the ER membranes are converted to large sacs, which can attach to the NE but fail to contribute to NE expansion. This suggests the possibility that the organisation of the interface between the NE and ER, as observed in oocytes, may be important for NE growth. Therefore we speculate that a possible function of localising Rtn4a to the inter-membrane junctions between ONM and ER is to maintain a particular interfacial organisation which contributes to the movement of membrane into the NE. Our results argue against a model in which membranes simply diffuse into the NE during assembly and growth (Ellenberg et al., 1997) because the NE fails to grow when Rtn4a is perturbed despite the attachment of ER membranes to the NE.
Rtn4a is not only located at the inter-membrane junctions but also throughout the tubular ER, where it appears to be involved in maintaining the tubularity (Voeltz et al. 2006; Fig. 7). Therefore, the tubular nature of the ER may be essential for its function in providing membrane for NE growth, at least in egg extracts.
The Rtn4a antibody does not inhibit the initial formation of NE around chromatin. This NE appears normal: it is flattened, fused and contains NPCs. Because it contains NPCs we can conclude that the contributing membranes carried the integral membrane nucleoporins such as POM121 and gp210 enabling NPC formation. This suggests that Rtn4a may only be required during the growth phase and that membranes that contribute to initial enclosure and NPC formation (Yang et al., 1997; Drummond et al., 1999) are not perturbed by the antibody.
We have also found that Rtn4a locates specifically to the edges of flattening membranes in growing and assembling NE in both in vivo and in vitro experiments. As Rtn4a has been implicated in the formation of highly curved membrane regions, it is possible to speculate that Rtn4a is important for formation of flattened sheets, by stabilisation of the high curvature at the edge regions. However, this may be a non essential function in the initial stages of NE assembly, which occur in the presence of the antibody.
4
Conclusions
We have shown for the first time that Rtn4a partitions, within a single membrane structure, to the region of highest curvature, supporting the model that reticulons preferentially locate to curved membranes (Voeltz et al., 2006) and have a function related to membrane curvature. We have also observed that it locates to membrane junctions and other highly curved regions of membranes that may be involved in NE growth. Concordantly, an Rtn4a antibody perturbs NE growth. We therefore hypothesise that Rtn4a may have a role in maintaining functional ER–NE junctions during NE growth and/or the ER must be tubular to contribute to NE growth.
4.1
Materials and methods
4.1.1
Isolation and fixation Xenopus oocyte for feiSEM
Stage III and VI NEs were isolated in 5:1 buffer, spread, fixed in 2% glutaraldehyde, 0.2% tannic acid, 10 mM Tris–HCl (pH 7.4) and 0.3 mM MgCl2 and processed for feiSEM as described previously (Kiseleva et al. 2004). In some experiments tannic acid, which helps preserve protein filament (Maupin and Pollard, 1983) was omitted, as it can effect membrane preservation (MWG unpublished), but the same membrane structures were observed. Samples were coated with chromium using a Cressington 308R with additional cryo-pump or an Edwards Auto306 with cryo-pump to a nominal thickness of 2 nm. They were viewed using a Hitachi S-5200 feiSEM at 10 kV accelerating voltage.
4.1.2
Fixation Xenopus oocyte for TEM
Oocytes were fixed (2.5% glutaraldehyde, 0,1M Hepes) for 1 h, then washed twice in 0.1 M Hepes buffer and postfixed 1 h in 1% OsO4 in ddH2O at 4 °C, stained 2 h in 1% aqueous uranyl acetate, washed in water, dehydrated through ethanol series and embedded in Agar-100 (Agar Scientific, UK). Sections were stained with Lead citrate and viewed with Leo 910 (Germany) TEM at 80 kV.
4.1.3
Immuno-TEM
Stage III oocytes were fixed in 4% formaldehyde (TAAB Labs) in Ringers (111 mM Nacl, 1.9 mM KCl, 1.1 mM CaCl2, 2.4 mM NaHCO3, pH 7.0) overnight at 4 °C, washed three times in Ringers, stain in 2% uranyl acetate in Ringers for 2 h at 4 °C, then washed in water twice, dehydrated in 30% ethanol, then further dehydrated by freeze substitution as follows through a series of ethanol (30% at 4 °C for 1 h, 50% at −20 °C for 1 h, 70% at −20 °C for 1 h, 95% at −20 °C for 2 h), embedded to LR Gold (Agar) at −20 °C and polymerised in gelatin capsules under UV light at −15 °C for 48 h. Ultrathin sections were cut and attached to nickel grids then incubated with 1% BSA in PBS for 30 min, washed in PBS, incubated with 1:100 dilution anti-Rtn4a rabbit antibody for 1 h, washed in PBS and incubated with a goat anti-rabbit secondary antibody conjugated to 10 nm colloidal gold (Amersham) for 1 h and washed with PBS. Sections were stained with lead citrate for 2 min.
4.1.4
Immuno-gold labelling and feiSEM imaging
Affinity purified polyclonal antibody (40 M stock) against N-terminal domain of Xenopus Rtn4a (Voeltz et al., 2006) was diluted 1:100 with PBS. Nuclei were isolated and fixed for 20 min in 3.7% formaldehyde, 5:1 buffer, washed three times with PBS, incubated in PBS, 1% BSA 30 min, washed in PBS, and incubated 1–3 h primary antibody, PBS. Samples were washed three times in PBS, and incubated 1 h with 10 nm gold-conjugated secondary goat anti-rabbit antibody (Amersham Corp.). As negative controls, we used gold-conjugated secondary antibody diluted 1:100 in PBS. All samples were then washed three times in PBS then in 10 mM Tris–HCl and processed for feiSEM as above. Both secondary electron images (for structure) and backscatter electron images (for gold label position) were collected simultaneously. Using Adobe Photoshop, the backscatter image was superimposed onto the secondary image as a separate layer. The positions of the gold particles were marked with a yellow dot and then the backscatter image removed.
4.1.5
In vitro nuclear assembly
Egg extracts were prepared as described previously (Goldberg et al., 1997). To perturb Rtn4a, extract was incubated with 4 μM affinity purified Xenopus Rtn4a antibody (Voeltz et al., 2006) on ice for 20 min then 1000 sperm chromatin per microlitre was added and the extract warmed to room temperature for 2–60 min for assembly. In controls an equal volume of either buffer or irrelevant antibody was added, which had no detectable effect on nuclear assembly. Extracts were resuspended in Membrane Wash Buffer (MWB: 250 mM sucrose, 50 mM KCl, 2.5 mM MgCl2, 10 mM Hepes, pH 7.4) and centrifuged at 1000g at 4 °C for 10 min onto silicon chips (Agar Scientific Ltd) and immersed in Membrane Fix (150 mM sucrose, 1 mM MgCl2, 80 mM Pipes–KOH, pH 6.8, 2% paraformaldehyde, 0.25% glutaraldehyde) for 10 min and processed for feiSEM as previously described (Goldberg et al., 1997). Samples were imaged at 3 kV accelerating voltage.
4.1.6
Antibody labelling of in vitro nuclei
Extract (5 μl) was resuspended in MWB and centrifuged at 1000g at 4 °C for 10 min onto silicon chips (Agar Scientific, UK). Chips were immersed in Membrane Fix without glutaraldehyde for 10 min, washed in PBS, immersed in PBS +100 mM glycine 10 min, washed in PBS, immersed in 1% fish skin gelatine (Sigma) for 1 h, then 1 μM primary antibody for 1 h, washed 3 times 5 min in PBS, incubated with 1:50 dilution anti-rabbit secondary antibody conjugated to 10 nm gold (Amersham) for 1 h, washed 3 times 5 min and 1 times 15 min in PBS and fixed in Membrane fix with 1% glutaraldehyde. Samples were then processed for feiSEM as previously described (Goldberg et al., 1997).

reticulon 4a
scanning electron microscopy
nogoa nuclear envelope


Muscle precursor cells in the developing limbs of two isopods (Crustacea, Peracarida): an immunohistochemical study using a novel monoclonal antibody against myosin heavy chain

In the hot debate on arthropod relationships, Crustaceans and the morphology of their appendages play a pivotal role. To gain new insights into how arthropod appendages evolved, developmental biologists recently have begun to examine the expression and function of Drosophila appendage genes in Crustaceans. However, cellular aspects of Crustacean limb development such as myogenesis are poorly understood in Crustaceans so that the interpretative context in which to analyse gene functions is still fragmentary. The goal of the present project was to analyse muscle development in Crustacean appendages, and to that end, monoclonal antibodies against arthropod muscle proteins were generated. One of these antibodies recognises certain isoforms of myosin heavy chain and strongly binds to muscle precursor cells in malacostracan Crustacea. We used this antibody to study myogenesis in two isopods, Porcellio scaber and Idotea balthica (Crustacea, Malacostraca, Peracarida), by immunohistochemistry. In these animals, muscles in the limbs originate from single muscle precursor cells, which subsequently grow to form multinucleated muscle precursors. The pattern of primordial muscles in the thoracic limbs was mapped, and results compared to muscle development in other Crustaceans and in insects.
Introduction
Amongst all arthropods, Crustaceans display the greatest variety of different limb types, and the morphology of their appendages has played an outstanding role in the raging debate on arthropod phylogeny (reviews, e.g. Williams and Nagy 1996; Boxshall 1997; Kukalova-Peck 1997; Browne and Patel 2000; Bitsch 2001; Klass and Kristensen 2001; Schram and Koenemann 2001; Williams and Nagy 2001; Wolf and Harzsch 2002; Waloszek 2003; Williams 2004). The recent move towards integrating the fields of evolutionary and developmental biology (e.g. Averof and Akam 1995; Gilbert et al. 1996; Akam 1998a; Hughes and Kaufman 2000; Harzsch and Hafner 2006; Harzsch 2007) has fostered a surge of studies on Crustacean limb development which examined the expression and function of genes such as Distal-less (Panganiban et al. 1995; Popadic et al. 1996, 1998; Scholtz et al. 1998; Williams 1998, 2008; Williams et al. 2002), Ultrabithorax and AbdominalA (Averof and Akam 1995; Averof and Patel 1997), Extradenticle (Gonzáles-Crespo and Morata 1996; Abzhanov and Kaufmann 2000), Pdm and Apterous (Averof and Cohen 1997), Sex combs reduced (Abzhanov and Kaufman 1999), and Wingless (Nulsen and Nagy 1999) in various Crustacean taxa with uniramous, biramous or phyllopodous branched limbs. Interestingly, some of these studies failed to establish homologies between the function of these genes during development of the complex Crustacean limbs as compared to the uniramous limbs of Insecta (Williams and Nagy 1995, 1996; Averof and Patel 1997; Williams et al. 2002; Williams 2004) but instead established new hypotheses on the evolution of hox gene function (Averof et al. 1996; Akam 1998b). The emerging picture is that limb patterning genes seem to act differently in the insect with uniramous limbs and those Crustaceans with phyllopodous limbs, and therefore, a greater knowledge of the cellular foundations of limb development in Crustaceans is essential to establish an interpretative context in which to analyse gene functions. However, few papers have recently dealt with cellular aspects of Crustacean limb development other than gene expression (e.g. Williams and Müller 1996; Ungerer and Wolff 2005; Kiernan and Herzler 2006).
Concerning the neuromuscular innervation, there is evidence for close similarities between Hexapoda and malacostracan Crustacea. In these animals, each thoracic walking leg is supplied by a set of exactly three inhibitory motoneurons in addition to its excitatory innervation. Wiens and Wolf (1993) have shown that the inhibitory limb innervation in a crayfish displays striking similarities to that in Hexapoda down to the level of single identified cells. The sets of inhibitors in these taxa share a number of morphological, physiological and biochemical characteristics which suggest homology, as discussed in greater detail by Harzsch (2007). Furthermore, the innervation pattern of particular excitatory motoneurons in crayfish and locusts provides new insights into the alignment of malacostracan Crustacean and insect trunk limbs (Wiens and Wolf 1993). These authors suggest a homology of the extensor muscles located within the second podomeres of insect and malacostracan limbs (merus and femur) and therefore support a close correspondence of limb segmentation in Malacostraca and Hexapoda (discussed in more detail by Wolf and Harzsch 2002; Harzsch 2007). Because information on inhibitory and excitatory leg motoneurons so far is only available for malacostracan Crustacea but not for the other Crustacean taxa, these comparisons so far are only of a limited phylogenetic value. However, these studies signify that comparative analyses of the neuromuscular system have a significant potential to contribute new insights into the evolution of arthropod appendages.
The cellular basis of embryonic muscle formation in Crustaceans is poorly understood, although ontogenetic aspects of the neuromuscular system (reviewed in Govind 1982; Govind and Walrond 1989; Govind 1995) and moult-induced muscle atrophy and regeneration (reviewed in El Haj 1999; Mellon 1999; Mykles 1999; Govind 2002) have been studied in some detail. The goal of the present project was to analyse muscle development in Crustacean appendages, and therefore, we generated monoclonal antibodies against arthropod muscle proteins. One of these antibodies, 016C6, strongly labelled muscle precursor cells in malacostracan Crustacea and on Western blots was shown to recognise several isoforms of myosin heavy chain in rabbit, grasshopper and Crustaceans. We used this antibody to study muscle formation in two isopods, the terrestrial Porcellio scaber Latreille, 1804 (Crustacea, Malacostraca, Peracarida, Isopoda, Oniscidea; Fig. 1) and the marine Idotea balthica Pallas 1772 (Valvifera) by immunohistochemistry. In these animals with uniramous walking legs (Fig. 1), muscles of the body wall and the limbs were shown to originate from single muscle precursor cells, which subsequently grow to form multinucleated muscle precursors. The pattern of primordial muscles in the thoracic limbs was mapped, and results compared to muscle development in insects (reviewed in Campos-Ortega and Hartenstein 1997; Abmayr and Keller 1998; Baylies et al. 1998; Paululat et al. 1999a, b; Roy and VijayRaghavan 1999).
Fig. 1The walking limbs of adult isopods: A ventral view of an adult male of I. balthica. The labels Th2 to Th8 identify the thoracomeres. B Adult specimen of P. scaber, ventral view (medial is towards the right) of thoracomeres three to eight (Th3–Th8) with the walking limbs attached. C Higher magnification of the walking limb from thoracomere three (P. scaber) to show podomeres (medial is towards the left). Abbreviations: letters in the upper right corners identify the species
Materials and methods
Animals
I. balthica Pallas 1772 (Crustacea, Malacostraca, Peracarida, Isopoda, Valvifera; Fig. 1A) were obtained from the Biologische Anstalt Helgoland and kept in artificial seawater at 16°C at the University of Konstanz (compare Kreissl et al. 1999). Eriphia spinifrons (Crustacea, Malacostraca, Decapoda, Brachyura) and Idotea emarginata (Crustacea, Malacostraca, Peracarida, Isopoda, Valvifera) were obtained from the Stazione Zoologica Anton Dhorn in Naples. P. scaber Latreille, 1804 (Crustacea, Malacostraca, Peracarida, Isopoda, Oniscidea; Fig. 1B, C) were collected on the campus surrounding the University of Konstanz. For the experiments, embryos were gently removed from the brood pouches of ovigerous females.
Generation and characterisation of the antibody 016C6
A crude myosin extract (d’Albis et al. 1979) of meso- and metathoracic locust muscles was used as the antigen to immunise mice. Two mice (female Balb/c, 6 weeks of age) received 0.1 ml of myosin extract emulsified in a 1:1 relation in RIBI adjuvant system, (MPL+TDM emulsion, R-700; RIBI Immunochem Research Inc.) at each of two subcutaneous sites on day 0 and day 28 and three final immunisations intraperitoneally with 50 μl myosin extract after 15 weeks. The mice sera were tested for antibodies on enzyme-linked immunosorbent assay (ELISA) coated with crude myosin extract. Both mice showed strong immune responses. Three days later, splenectomy was performed, and the splenocytes were fused with a mouse myeloma cell line (P3X63Ag8.653, ATCC CRL-1580) by PEG-1500. The successfully fused hybridomas were selected in Roswell Park Memorial Institute (RPMI)-1640 medium supplemented with HAT (10 mM hypoxanthine, 40 μM aminopterin, 1.6 mM thymidine; GIBCO BRL, Carlsbad, CA, USA), 10% foetal bovine serum, 24 μM β-mercaptoethanol and Nutridoma-CS (Roche, Indianapolis, IN, USA). The culture supernatants were collected and screened by ELISA. The reactions were confirmed with Western blot analysis and immunostaining on cryosections of native and paraformaldehyde fixed locust muscles using secondary antibody conjugated to Cy3 or Cy2 (Jackson Labs). Positive clones were selected by at least three sets of the limiting dilution technique. The antibodies generated against locust muscle proteins were also screened for their binding affinity to muscle tissue of malacostracan Crustaceans and of rabbits. For the identification of the antigen in Western blots, the crude myosin extracts (d’Albis et al. 1979) of locust muscles, of abdominal muscles of the two Crustacean species E. spinifrons and I. emarginata and of adult White New Zealand rabbit muscles (psoas, soleus and diaphragm) were separated by sodium dodecyl sulfate–polyacrylamide gel electrophoresis (SDS-PAGE) on 7.5% linear gels according to standard techniques. The proteins were transferred to cellulose nitrate membranes (Protran, Schleicher & Schuell GmbH, Dassel, Germany), and selective binding of monoclonal IgG1κ 016C6 to proteins was detected with the IgG-ABC-ELITE-POD kit (Vector Labs) using ECL (Pierce) as a substrate. Molecular weight markers were biotinylated, allowing direct detection by the ABC reagent in the Western blots.
Whole mount immunohistochemistry
Embryos were fixed for 4 h in 4% paraformaldehyde in 0.1 M phosphate buffer (pH 7.4) at room temperature. Whole mounts of the embryos were incubated in 1 mg/ml collagenase/dispase (Sigma) for 20 min at room temperature or pretreated with short ultrasonic pulses to increase penetration of the antibodies then washed in several changes of 0.1 M phosphate-buffered saline (PBS) for 4 h and afterwards pre-incubated in PBS containing 1% normal goat serum and 0.3% Triton X-100 (PBS-TX) for 2 h at room temperature. Specimens were then incubated overnight at 4°C in supernatants of the monoclonal antibody 016C6 diluted 1:10 in PBS-TX. The omission of the primary antibody resulted in a complete absence of specific labelling. The embryos were then incubated in a biotinylated secondary antibody for 3 h (Jackson) and subsequently for another 3 h in peroxidase-conjugated streptavidin (Dianova). After washing for 4 h in PBS, the tissues were reacted with 0.013% diaminobenzidine and a reagent containing hydrogen peroxide, cobalt chloride and nickel chloride (Amersham, RPN 20) for 7–9 min to reveal the peroxidase label. Finally, whole-mount preparations were dehydrated and mounted in Eukitt (Riedel-de Haen). Preparations were observed with a Zeiss Axioskop and labelled structures drawn using a camera-lucida apparatus. The slides were also photographed on 35-mm colour slide film, the images transferred onto Kodak Photo CD and processed in Picture Publisher. Alternatively, specimens were photographed with the Polaroid DMC10 digital camera.
HOECHST stain
To reveal the morphology of the early embryos, specimens were dissected out of the chorion and the yolk was removed. Specimens were then fixed in 4% paraformaldehyde in 0.1 M phosphate buffer (1 h, room temperature) and stained with the nuclear dye bisbenzimide (0.1%, 15 min at room temperature; Hoechst H 33258), washed in buffer overnight and mounted in Flouromount (Sigma). Specimens were viewed with a fluorescent microscope (Axioskop) and documented as described above.
Results
Characterisation of monoclonal antibody 016C6
Screening the supernatants of a number of monoclonal hybridoma cell lines producing antibodies against locust muscle proteins for their binding affinity to muscle tissue of other species revealed that monoclonal antibody (MAB) 016C6 shows a particularly strong affinity for muscle proteins of malacostracan Crustaceans. For the identification of the 016C6 antigen, muscle proteins of Locusta migratoria, I. emarginata, E. spinifrons and of adult White New Zealand rabbits were separated by SDS-PAGE and analysed in Western blots. Polyacrylamide gel analysis of proteins of the muscle homogenates shows prominent bands with apparent molecular weights of 180 to 200 kDA, indicating the presence of myosin heavy chains in all tissues examined (Fig. 2A). In the four tested species, MAB 016C6 binds to muscle proteins with an apparent molecular weight of 180 kDA (Fig. 2B).
Fig. 2Characterization of monoclonal antibody 016C6: 016C6 specifically binds to myosin heavy chains, which co-migrates with a slow myosin heavy chain isoform of rabbit muscles. A Separation of muscle proteins by SDS-PAGE. Numbers at the left indicate molecular weight markers (MW), lanes 2–4 were loaded with muscle homogenates of the two malacostracan species E. spinifrons (Decapoda, Brachyura) and I. emarginata (Peracarida, Isopoda) and the hexapod L. migratoria. Lanes 5–7 were loaded with homogenates of the fast type psoas muscle, the mixed type diaphragm and the slow type soleus muscle of rabbits. Myosin heavy chains are prominent at MW of 180 to 200 kDA. B Identification of the 016C6 antigen by Western blot analysis reveals myosin heavy chains in arthropod muscles which co-migrate with a 180 kDA myosin heavy chain isoform of rabbit slow and mixed muscles
In mammals, myosin heavy chains (MHC) exist as isoforms of polypeptides with a molecular mass of about 180–200 kDa. The homogenate of the rabbit psoas (fast twitch muscle) contains predominantly the MHC-IId isoform. The homogenate of the diaphragm contains MHC I, MHC IIa and MHC IId, the slow twitch and two fast twitch isoforms, respectively. The rabbit soleus (slow twitch muscle) contains predominantly myosin heavy chain MHC-I (Aigner et al. 1993). MAB 016C6 exhibits a considerably higher affinity to MHCs of the three arthropod species and to the slow MHC-I isorm of rabbit muscles as compared to the fast rabbit MHCs (Fig. 2). We used MAB 016C6 as a general marker for differentiating muscle cells already containing myosin heavy chains in the present study.
Development of the embryos
The embryonic development of two species of the genus Idotea has been described by Strömberg (1965), and the development of P. scaber was recently reviewed by Whitington et al. (1993) and Abzhanov and Kaufman (1999; see also Brena et al. 2005). Hejnol et al. (2006) explored germ band formation in this organism. For P. scaber, Whitington et al. (1993) established a percentage staging system based on developmental time. Under the rearing conditions described above, I. balthica embryos hatched after 27 days. However, a period of rapid organogenesis occurs during the last 6 or 7 days of embryogenesis (Fig. 3) so that a direct comparison of the ontogeny of I. balthica and P. scaber based on a percentage scale was not possible. Therefore, we subdivided the embryonic period that we studied into five distinct stages, which are comparable between the two species:
Stage 1(I. balthica E70%, P. scaber E40%; the percentage values indicate embryonic development in a percentage staging system based on developmental time): The germ band is not completely elongated, and mitotic cells in the posterior growth zone are still visible; in the more anterior thoracomeres, distinct limb anlagen can be distinguished.Stage 2(I. balthica E80%, P. scaber E60%): All segments are formed, the embryo is not closed dorsally, and the limb anlagen are subdivided into distinct podomeres.Stage 3(I. balthica E85%, P. scaber E80%): Red pigments is visible throughout the entire embryo, the first ommatidia can be distinguished, and single muscle precursor cells can be labelled immunohistochemically in the limb anlagen.Stage 4(I. balthica E90%, P. scaber E90%): The growing tergites have dorsally enclosed the yolk.Stage 5(I. balthica E100%, P. scaber E100%): hatchingFig. 3Embryonic staging system: A–C embryos of I. balthica at A 70% of embryonic development (E70%; stage 1), B E85% (stage 3) and C E95% (stage 4–5). D Embryo of P. scaber at E60% (stage 2). E, F embryos of P. scaber at E E40% (stage 1) and F E60% (stage 2), HOECHST nuclear stain, ventral views. At stage 1, the germ band is not completely elongated, and limb anlagen are visible in the anterior thoracic segments. At stage 2, all segments are formed, and the thoracic limb anlagen are subdivided into distinct podomeres. Abbreviations: letters in the upper right corners identify the species and larval stages; A1-2 antenna 1 and 2, GZ posterior growth zone, LB labrum, MD mandible, MX1-2 maxilla 1 and 2, T1-8 thoracomeres 1 to 8, P1-5 pereomeres 1 to 5, the last pleomeres denoted. Scale bars A–F, 100 μm
Development of the thoracic limbs
A stage 1, embryo of P. scaber labelled with the HOECHST stain is shown in Fig. 3E. Proliferating cells in the posterior growth zone have just generated the first pleomere, and limb buds are beginning to form in the thoracic segments at that stage. The thoracic limb anlagen subsequently enlarge, and bilobed anlagen of the pleopods appear (Fig. 4A, late stage 1). In stage 2 embryos, the developing thoracopods begin to subdivide into distinct subunits, the podomeres (Fig. 3E, F4B), and the adult subdivision of the thoracopods into seven subunits is established: coxa, basis and the five endopodal podomeres ischium, merus, carpus, propodus, dactylus (Fig. 4B). At this stage, the limb anlagen are well developed in all pleonic and thoracic segments except thoracomere 8. The eighth thoracopods are not present in embryos (Fig. 4A,B) but develop postembryonically (Fig. 1; see Abzhanov and Kaufman 1999).
Fig. 4Development of the walking limbs: A late stage 1 embryo (P. scaber), limb anlagen are present in the thoracic and pleonic segments except thoracomere 8; Normarsky interference contrast, ventral view, anterior is towards the right in all images of this figure. B stage 2 embryo (P. scaber), the thoracic limbs are subdivided into characteristic podomeres, thoracopod 8 is still undeveloped, lateral view, HOECHST nuclear stain. C–E Immunohistochemistry with monoclonal antibody 016C6 (I. balthica). Muscle precursor cells in thoracic limbs 5 and 6 (C), antenna 2 of stage 3 embryos (D) and matured muscle in thoracopod 5 of a stage 5 embryo (E). Abbreviations: letters in the upper right corners identify the species and larval stages; Ba basis, Ca carpus, CX2-7 coxae of thoracopods 2 to 7, Da dactylus, Is ischium, Me merus, P1-5 pleomeres 1 to 5, Pr propodus, T8 thoracomere 8. Scale bars: A 30 μm, B 50 μm, C–E 30 μm
Immunohistochemistry with monoclonal antibody 016C6 revealed that muscle mononucleate precursor cells that express myosin heavy chain can be labelled for the first time in stage 3 embryos of I. balthica and P. scaber (Fig. 4A,C). Sets of muscle precursors are arranged in a repetitive pattern within each podomere of the thoracic limbs and also the cephalic appendages, e.g. the second antennae (Fig. 4C,D). During subsequent development, these precursors enlarge to form a complex pattern of muscle precursors in the thoracopods (Figs. 4E, 5, 6 and 7).
Fig. 5Myogenesis in the limb anlagen: Immunohistochemistry with monoclonal antibody 016C6 in the propodus of the thoracic limbs. Muscles propodus one and two (Pr1, Pr 2) originated from single, mononucleate cells, the muscle precursors or muscle founder cells, both in stage 3 embryos of P. scaber (A) and I. balthica (B). During subsequent development, the precursor cell of Pr1 enlarged (C) and eventually developed into a binucleate, syncytial muscle precursor (D, I. balthica). This muscle precursor further enlarged in size (E) and became a multinucleate muscle precursor (F, G, I. balthica). In stage 4 embryos, both muscles, Pr1 and the antagonistic Pr2, had developed into substantial primordial muscles (H). In stage 5 embryos, before hatching, Pr1 and Pr2 had subdivided into two or three distinct subunits (I, I. balthica). J: Development of muscles Pr1 and Pr2 in the propodus of I. balthica, schematic representation. Abbreviations: letters in the upper right corners identify the species and larval stages; Ca carpus, Da dactylus, Pr propodus. Scale bars: A–I 5 μm (scale bar in B valid also for C–F)Fig. 6Muscles in the whole mount of a pre-hatching embryo: Immunohistochemistry with monoclonal antibody 016C6, stage 5 (pre-hatching) embryo of I. balthica. Abbreviations: A1, 2 antenna 1 and 2, MD mandible, MX1, 2 maxilla 1 and 2, OP operculum, P1-3 pleopods 1 to 4, T1-7 thoracic limbs 1 to 7 (T1 is a maxilliped), T8 thoracomere 8. Abbreviations: letters in the upper right corners identify the species and larval stages. Scale bar 100 μmFig. 7Map of muscle precursor in a walking limb: A Immunohistochemistry with monoclonal antibody 016C6, thoracic limb 5, stage 5 (pre-hatching) embryo of I. balthica, frontal view, medial is towards the left. B Schematic drawing of the muscles in thoracic limb 5 of a stage 5 (pre-hatching) embryo of I. balthica (frontal view as in A). Anteriorly arranged muscles are drawn in light grey, posteriorly arranged muscles are drawn in dark gray. The color scheme of the podomeres is according to Walossek (1999) and Waloszek (2003); see also Wolf and Harzsch (2002): coxa—red, basis—yellow, endopodite—green. C Schematic representations of the directions of movement of the joints between the podomeres. Abbreviations: letters in the upper right corners identify the species and larval stages; Ba basis, Ca carpus, Da dactylus, Is ischium, Me merus, Pr propodus; for the terminology to label the muscles see text. Scale bar 25 μm
Formation of a single identified muscle in the propodus of the thoracic limbs
Myogenesis will be exemplified by the formation of the identified, antagonistic muscles propodus 1 and 2 (Pr1, Pr2; Fig. 5), both of which are located in the propodus of the thoracic limbs and which move the dactylus in the adult (see below). Pr1 originates from single, mononucleate muscle precursors or muscle founder cells, which express myosin heavy chain both in stage 3 embryos of P. scaber (Fig. 5A,J; the mononucleate precursor cell of muscle Pr2 is also shown) and I. balthica (Fig. 5B). During subsequent ontogeny, this mononucleate precursor cell enlarge (Fig. 5C) and eventually develop into a binucleate, syncytial muscle precursor (Fig. 5D,J). This muscle precursor further enlarges in size (Fig. 5E) and becomes a multinucleate muscle precursor (Fig. 5F,G,J). In stage 4 embryos, both muscles, Pr1 and the antagonistic Pr2, have developed into substantial primordial muscles (Fig. 5H). In stage 5 embryos, before hatching, Pr1 and Pr2 have subdivided into two or three distinct subunits (Fig. 5I,J).
Muscle pattern in the thoracic limbs of pre-hatching embryos
Immunohistochemistry with 016C6 enabled us to map the primordial muscles in the thoracic appendages of I. balthica embryos before hatching (Figs. 6 and 7). As the adult pattern of muscles has not been mapped in the isopods which we examined, we labelled the primordial muscles with two letters to indicate the podomere in which they were located plus a number counting up the muscles from anterior to posterior (Fig. 7B). Me2, for example, designates muscle number 2 in the merus. Most embryonic muscles are arranged in antagonistic groups and are restricted to one podomere (Fig. 7B). Exceptions are Ba/Is1 and Ba/Is2, which span across the basis and the ischium (Fig. 7B). In the coxae, a complex system of muscles is present, which we failed to map.
To determine the function that the embryonic muscles will exert in the adult limbs, we examined the articulations and directions of movement of the successive podomeres in adult limbs of P. scaber (Fig. 1B,C) and I. balthica (Fig. 1A). These functions can be tentatively summarised as follows (Fig. 7B,C, Table 1): Ba7 serves as abductor of the ischium while Ba1, 2, 3 and Ba4, 5 serve as adductors of the ischium that swings in a medial to lateral plane. The merus also moves in a medial to lateral plane and is abducted by Is1 and Ba/Is1 and adducted by Ba/Is2 and Is 2, 3. The carpus swings in an anterior to posterior plane perpendicular to the merus. It is retracted by Me2 and protracted by Me1. The carpus also seems to have a limited freedom of movement in the medial to lateral. The propodus once more is displaced exclusively in a medial to lateral plane with Ca3 serving as the abductor and Ca1 and Ca2 as adductors. The dactylus swings in the same plane and is adducted by Pr1 and abducted by Pr2 and Pr3. The coax–basis joint was not examined in detail but seemed to allow displacement of the basis into several directions (Alexander 1972).
Table 1Muscles in the walking limbs of crayfish and other Decapoda (Evoy and Ayers 1982; Atwood and Govind 1982; “common” names are also indicated), Isopoda (present report and Alexander 1972), Amphipoda (Brusca 1981)Muscles located in this podomereCrayfish and other DecapodaIdotea (Isopoda)Cystisoma (Amphipoda)ThoraxNot shownNot shownNot shownCoxa2 Depressors of basis2 Extensor of basis2 Retractors of basis2 Levators of basis2 Flexors of basis2 Protractors of basis2 Rotators of basisadductor of basisBasisFlexor of ischium2 Flexors of ischiumBasis and ischium are fused in adult Decapoda and contain the reductor of merus3 Lateral extensors of ischium2 Extensors of ischium2 Lateral extensors of ischiumIschium2 Flexors of merusFlexor of merus2 Lateral extensors of merus2 Extensors of merus2 medial extensors of merusMerusAbductor of carpus (extensor)Abductor of carpus3 Abductors of carpusAdductor of carpus (flexor)Adductor of carpus2 Adductors of carpusAccessory adductor of carpusCarpusReductor of propodus (stretcher)Flexor of propodus2 Flexors of propodusProductor of propodus (bender)Lateral extensor of propodus2 Extensors of propodusAdductor of propodus (rotator)Medial extensor of propodusPropodusAbductor of dactylus (opener)Flexor of dactylus3 Flexors of dactylusAdductor of dactylus (closer)Lateral extensor of dactylus1 Extensor of dactylusmedial extensor of dactylusDactylusNo musclesNo musclesNo musclesMuscles located within the thorax are not shown.
Development of muscles in the body wall
Although muscle precursor cells are not present in the anlagen of the thoracic limbs in late stage 1 embryos (Fig. 4A), there is already a distinct pattern of muscle precursors in the body wall [Electronic supplementary material (ESM) Fig. 1]. A bilateral band of ventral longitudinal (VL) muscle precursors is arranged parallel to the midline (ESM Fig. 1A). This band displays an anterior–posterior of maturation so that several steps of muscle formation can be observed in a single specimen (ESM Fig. 1B,C). As in the limbs, the ventral longitudinal muscles also originate from single muscle mononucleate precursor cells (inset ESM Fig. 1A), which span the length of one segment. These precursors subsequently develop into bi- (ESM Fig. 1B) and multinucleated muscle precursors (ESM Fig. 1C). In late stage 1 embryos, a band of dorsal longitudinal (DL) muscles is arranged in parallel to the ventral longitudinal muscles (ESM Fig. 1D,E). The ventral and dorsal bands of muscles are connected by single, segmentally iterated transverse muscle precursor cells (arrows in ESM Fig. 1E). In later stages, 016C6 also labelled a dense network of muscles surrounding the hindgut (ESM Fig. 1F).
During subsequent ontogeny, the muscle precursors of the longitudinal muscles mature into a more complex system of dorsal (extensors, ESM Fig. 2A) and ventral (flexors, ESM Fig. 2C) body wall musculature. A comparison of muscles in the leg-bearing thoracic segment and the leg-less eighth thoracomere enabled us to distinguish between muscles of the body wall and those that are associated with the coxa (Fig. 6, ESM Fig. 2B). In addition to the ventral and dorsal longitudinal muscles, distinct intersegmental extensor muscles are present in the eighth and the more anterior thoracomeres. These muscles transverse the border between two successive thoracomeres (Fig. 6, ESM Fig. 2A,B). Furthermore, two smaller transverse muscles are found in each thoracomere (ESM Fig. 2B).
Discussion
Myogenesis in Crustacea
In recent years, cell lineages analyses in malacostracan embryos have set out to explore the earliest stages of mesoderm formation (Gerberding et al. 2002; Hertzler 2002, 2005), and the molecular mechanisms that underlie mesoderm formation have been explored in the emerging Crustacean model system, the amphipod Parhyale hawaiensis (Price and Patel 2008). Furthermore, in decapod Crustaceans, considerable effort has been directed towards understanding aspects of the ontogeny of the neuromuscular system such as outgrowth of motoraxons, development of the innervation patterns of muscles and synapse formation. These processes have been examined, e.g. in the body wall of the pleon (Cole and Lang 1980; Stephens and Govind 1981; Govind et al. 1985), the pleopods (Davis and Davis 1973; Kirk and Govind 1992) and the thoracic limbs (Costello et al. 1981; Govind and Pearce 1981, 1982, 1989; Govind et al. 1982; Govind and Derosa 1983; Pearce et al. 1985; Lnenicka et al. 1991; Arcaro and Lnenicka 1995). Myogenesis has been studied in the pleon of embryonic American lobsters on the electron-microscopic level (review Govind 1982, 1995). According to these accounts, the first signs of muscle formation are the appearance of small localised patches of myofilaments in multinucleated structures, the presumptive myotubes. Surrounding undifferentiated cells have enlarged nuclei with diffuse chromatin, Golgi apparatus, ribosomes and mitochondria and seem to resemble premyoblast cells (Govind 1982, 1995). The myofilaments then become organised in longitudinal arrays, and later, distinct sarcomeres with A, Z and I bands appear. The muscle fibres grow in length either by the elongation of individual sarcomeres or the serial addition of sarcomeres (Govind 1982, 1995). Muscle development has also been studied in the embryos and nauplii of the dendrobranchiate shrimp Sicyonia ingentis using fluorescent phallotoxins to label F-actin (Kiernan and Hertzler 2006). In these animals, phalloidin labelling identifies muscle precursor that in the beginning are not striated and during mid-embryogenesis stretch along the entire length of the naupliar appendages (antenna one, antenna two and mandible). In this study, the analysis of myogenesis was restricted to these naupliar appendages and was followed up to the nauplius V stage when trunk appendages are still not developed (Kiernan and Hertzler 2006). Contrary to the dendrobranchiate S. ingentis which develops via nauplius larvae, the isopod species studied in the present study are direct developers so that a meaningful comparison of myogenesis in the trunk limbs is as yet not possible.
However, we can compare myogenesis in the pair of second antennae of this dendrobranchiate shrimp to the Isopoda. One major difference is that in Dendrobranchiata, the muscle precursors in antenna 2 span across the entire length of these appendages (Kiernan and Hertzler 2006), whereas in Isopoda, sets of individual muscle precursors are arranged in an iterated pattern within each podomere of the second antennae. Studies on appendage development in embryos of the American lobster Homarus americanus (Malacostraca, Homarida) using monoclonal antibody 016C6 (Harzsch and Kreissl, unpublished data) revealed that the pattern of myogenesis in this organism is different from that in Isopoda but rather resembles that in Dendrobranchiata. In lobster embryos, syncytial muscle precursor cells establish the muscles in the endopodites of the thoracic appendages and also in the second antennae. As in Dendrobranchiata, these muscle precursors initially stretch along the entire length of the appendages. During subsequent embryogenesis, the muscle precursors subdivide into several distinct units, thereby giving rise to iterated pairs of antagonistic primordial muscles in each of the successive podomeres (Harzsch and Kreissl, unpublished data), a layout which more and more resembles the arrangement in the adult thoracopods (see below and Table 1). We would predict that also in Dendrobranchiata, such an ontogenetic subdivision of the muscle precursors into individual iterated muscle primordia may take place, a question that should be explored by analysing later larval stages than have been studied so far. Considering that Kiernan and Hertzler (2006) reported striking similarities between the pattern of myogenesis between the dendrobranchiate S. ingentis and the brine shrimp Artemia salina (Crustacea, Branchiopoda) and considering that myogenesis in the American lobster H. americanus matches the Sicyonia/Artemia pattern (Harzsch and Kreissl, unpublished data), we conclude that these organisms most likely represent the ancestral Crustacean mode of myogenesis and that the slightly different mode that we observed in the present study for the Isopoda is derived from this ancestral pattern.
Comparison of myogenesis in Crustacea and Insecta
In the developing body wall of grasshopper embryos, Ho et al. (1983) were the first to describe large mononucleate mesoderm cells that arise early in development to erect a scaffold for later developing muscles, and they termed these cells muscle pioneers. The development of muscle pioneers in the body wall subsequently was also examined with antibodies different from that which Ho et al. (1983) used (Xie et al. 1992, 1994; Steffens et al. 1995). Furthermore, the role of muscle pioneers was extensively studied in the developing limbs of the grasshopper (Ball and Goodman 1985a, b; Ball et al. 1985) and a moth (Consoulas and Levine 1997; Consoulas et al. 1997). Since the discovery of muscle pioneers in the grasshopper, detailed analyses of myogenesis in the fruit fly has led to the establishment of the founder cell model for muscle patterning (reviewed in Campos-Ortega and Hartenstein 1997; Abmayr and Keller 1998; Baylies et al. 1998; Paululat et al. 1999a, b; Roy and VijayRaghavan 1999). According to this model, mesodermal progenitor cells in insects undergo mitosis to produce mononucleate muscle pioneers (termed founder cells in the fruit fly). These pioneers then differentiate into bi-, tri- or multinucleate syncytial muscle precursors by fusion with surrounding undifferentiated myoblasts.
Our present report applying a monoclonal antibody against myosin heavy chain extends the ultrastructural findings of Govind (1982, 1995) on muscle formation in malacostracan Crustaceans and suggests that in Isopoda, similar mechanisms of myogenesis as in insects may be present. This antibody labelled mononucleate myosin-expressing cells with morphological characteristics of insect muscle pioneers. Despite the differences between myogenesis in isopods and the American lobster noted above, the latter organisms nevertheless also seem to employ mononucleate myosin-expressing cells to found their limb muscles (Harzsch and Kreissl, unpublished data). Similar to insects, the muscle precursors in the limb anlagen of the isopod Crustaceans we examined were individually identifiable and established the primordial adult muscle pattern. Furthermore, these cells developed into bi- and multinucleate syncyctia similar to insect muscle precursors. However, our methods did not enable us to determine whether this process is achieved via fusion with surrounding myoblasts as it is in insects. We conclude that in the moment, our knowledge on Crustacean myogenesis is too limited and too few species studied as to suggest a homology of the mononucleate Crustacean muscle precursors and insect muscle pioneer cells. Although the musculature in the limbs of non-malacostracan Crustaceans has been examined in depth, e.g. in Copepoda (Boxshall 1985, 1990, 1997), Cephalocarida (Hessler 1964), Branchiopoda (Benesch 1969; Fryers 1988) and Cirripedia (Wallay 1969), myogenesis has only been studied in two representative of the Branchiopoda (Williams and Müller 1996; Kiernan and Hertzler 2006) and a cirripede (Semmler et al. 2006). However, these studies were not conducted at the cellular level that would allow for a meaningful comparison with our data. Therefore, it remains unclear at this point whether mononucleate muscle precursor cells are part of the Crustacean ground pattern. Similarly, as the mechanisms of muscle formation are not known in any representatives of the Chilopoda and Progoneata (Myriapoda), we cannot answer the question whether mononucleate muscle pioneers are a class of muscle founder cells that represent a synapomorphy of Hexapoda and Malacostraca or if they were already present in the ground pattern of Euarthropoda.
Muscle arrangement in the adult limbs of Isopoda and Decapoda
The structural organisation of the neuromuscular system in adult Crustacean limbs is particularly well understood in Decapoda (reviewed in Wiersma 1961; Evoy and Ayers 1982; Govind and Atwood 1982; Rathmayer and Maier 1986; Wiens 1989; Rathmayer 1990, 2002; Cattaert and Le Ray 2001; Clarac 2002). In this taxon, muscles in the limbs exert their force on a system of levers formed by apodemes and joints, most of which operate in a single plane. Usually, the movements of an individual joint are brought about by a pair of antagonistic muscles located in the podomere proximal to the podomere which they displace. Successive joints along a particular appendage operate at an angle of approximately 90° to one another, and most movements result from combinations of displacements at several joints. Three main joints are involved in locomotion: the thoraco-coxopodite joint allows forward and backward movements of the leg; the coxo-basipodite joint is responsible for upward and downward movements; the mero-carpopodite joint is responsible for extension and flexion of the leg (reviewed in Wiersma 1961; Evoy and Ayers 1982; Govind and Atwood 1982; Cattaert and Le Ray 2001; Clarac 2002).
In adult Isopoda, muscles have been extensively mapped in the cephalic appendages (Schmalfuss 1974) and the pleopods (Erhard 1997), but little information is available about the musculature of the thoracic limbs in the isopod taxa Oniscoidea and Valvifera. Von Haffner (1937) and Gruner (1954) described the morphology of the limbs in several isopod taxa but failed to individually identify the muscles, while Alexander (1972) only described the muscles located in the coxa. However, the limb musculature was mapped in several species of the amphipod genus Cystisoma Guérin-Méneville, 1842 (Hyperiidea; Brusca 1981). Table 1 summarises the muscle arrangement in adult Decapoda (Eucarida) and compares it with Amphipoda (Peracarida; Brusca 1981) and the isopod pre-hatching embryos (Peracarida) we studied. In all three species, the principle of paired antagonistic muscles (or muscle groups) is realised. However, while in Decapoda, in most cases, pairs of single antagonistic muscles are present, pairs of functional groups composed of two to five distinct muscles are found in Peracarida (von Haffner 1937; Brusca 1981). Furthermore, all successive joints in Decapoda operate at an angle of approximately 90° (Evoy and Ayers 1982), while in Peracarida, only the planes of the merus–carpus joint and the carpus–propodus joint are twisted against each other (Fig. 7; and von Haffner 1937; Brusca 1981). Another difference is that in adult Decapoda, basis and ischium are fused and bear only one muscle, the reductor of the merus. These two podomeres are unfused in Peracarida and therefore contain sets of flexors and extensors (Table 1; and von Haffner 1937; Brusca 1981).
Electronic Supplementary Material
Below is the link to the electronic supplementary material.
Supplementary Fig. 1
Myogenesis in the body wall: Immunohistochemistry with monoclonal antibody 016C6, muscle precursors in the body wall of late stage 1 embryos of P. scaber, dorsal views, anterior is towards the top. A bilateral bands of VL muscle precursors are arranged parallel to the midline. B, C Higher magnification of A, several steps of muscle formation from single muscle precursor cells (B and inset in A) to bi- and multinucleated muscle precursors (C) could be observed in this specimen. D, D′ A dense network of muscles surrounding the hindgut was labelled in later embryonic stages. E, F late stage 1 embryo, a band of DL muscles was arranged in parallel to the VL muscles. The ventral and dorsal bands of muscles were connected by single, segmentally iterated transverse muscle precursor cells (arrows, E). Abbreviation: T 1-7 thoracomeres 1 to 7. Scale bars: A, B 40 μm, C 20 μm, D 40 μm, D′ 10 μm, E 5 μm (F same magnification as E). (GIF 693 kb)
High resolution image file (TIF 15.0 MB)
Supplementary Fig. 2
Body wall muscles in a pre-hatching embryo: Immunohistochemistry with monoclonal antibody 016C6, mature system of muscles in the body wall of late stage 4 embryos of I. balthica, lateral views (A, C). The segmentally iterated dorsal longitudinal muscles (A) and ventral longitudinal muscles are visible (C), and the muscles that move the coxa can be discerned (asterisks). B A comparison of muscles in the leg-bearing thoracomere 7 segment and the leg-less eighth thoracomere enabled us to distinguish between muscles of the body wall (IE intersegmental extensors, TM transverse muscles; compare A) and those that are associated with the coxa (asterisks). Abbreviations: CX coxa, DL dorsal longitudinal muscles, extensors, IE intersegmental extensors, T7 thoracic leg 7, T8 thoracomere 8, VL ventral longitudinal muscles, flexors. Scale bars: A–C 30 μm. (GIF 650 kb)
High resolution image file (TIF 14.8 MB)

muscle precursor
crustacea
appendage
evolution
isopoda