Investigation of Hox gene expression in the Brazilian Whiteknee tarantula Acanthoscurria geniculata

Dan Strömbäck

Degree project in biology, Bachelor of science, 2020 Examensarbete i biologi 15 hp till kandidatexamen, 2020 Biology Education Centre and Institutionen för biologisk grundutbildning vid Uppsala universitet, Uppsala University Supervisor: Ralf Janssen

Abstract

Acanthoscurria geniculata, the Brazilian whiteknee tarantula, is part of the group Mygalomorphae (mygalomorph spiders). Mygalomorphae and Araneomorphae (true spiders) and Mesothelae (segmented spiders) make up Araneae (all spiders). All spiders have a prosoma with a pair of chelicerae, pedipalps and four pairs of legs, and an opisthosoma with two pairs of book lungs or one pair of book lungs and one pair of trachea (in opisthosomal segments O2 and O3) and one or two pairs of spinnerets (in segments O4 and O5). The mygalomorphs have retained two pairs of book lungs, an ancestral trait evident from looking at Mesothelae, the ancestral sister group of both Araneomorphae and Mygalomorphae. The spinnerets differ greatly between the groups, but this study focuses on comparing Mygalomorphae and Araneomorphae. Mygalomorphae also have reduced anterior spinnerets, but instead enormous posterior spinnerets. Araneomorphae possess all four, but none particularly big. The genetic basis of these differences between the set of opisthosomal appendages in tarantulas and true spiders is unclear. One group of genes that could be involved in the development of these differences could be the famous Hox genes. Hox genes have homeotic functions. If they are expressed differently between these two groups, the resulting morphology could change. This study focuses on the posterior Hox genes in A. geniculata, i.e. Antennapedia, Ultrabithorax, AbdominalA, and AbdominalB, each present in the form of two paralogs. Differences found in the expression patterns among these three spiders can explain structural differences in the opisthosoma.

Introduction

The Brazilian whiteknee tarantula, Acanthoscurria geniculata, belongs to the infraorder Mygalomorphae, a sister group of Araneomorphae (true spiders). They, along with the ancestral sister group Mesothelae (segmented spiders), make up the order Araneae. All spiders have some unique physical traits in common, such as the spinnerets, which are organs used to spin silk.

There are morphological differences between Mygalomorphae and Araneomorphae. Mygalomorphae are often much larger, and their chelicerates are angled differently from those in Araneomorphae. Mygalomorphae can only move their fangs vertically, whereas the fangs in Araneomorphae are angled toward each other. Mygalomorphae have also retained two pairs of book lungs, whereas in true spiders the posterior pair has been reduced to tracheae (Wheeler et al., 2016). Two pairs of book lungs are likely ancestral as the ancestral group Mesothelae also has two pairs (Küntzel et al., 2019). Other differences concern the spinnerets. Normally there are two pairs in the opisthosoma, two anterior in segment O4 and two posterior in segment O5. In Mygalomorphae, the anterior pair is reduced, but the posterior spinnerets are very large compared to those of Araneomorphae (Mariano-Martins et al., 2020; Wheeler et al., 2016).

These morphological differences could be explained by the function of the famous Hox genes. Hox genes are genetic “switches'' mainly active in the embryonic stage that turn on or off

downstream cascades that determine the identity of most segments (such as leg- , chelicerate- or spinneret-bearing segments). Their location on the chromosomes is in clusters, meaning that they are all next to one another (Pace et al., 2016). Interestingly, they are expressed in order of occurrence on the chromosome, and this is called co-linearity. The first Hox gene is activated earliest and is expressed most anteriorly in the embryo, the other Hox genes are then expressed one after one more and more posteriorly (Barnett & Thomas, 2013; Khadjeh et al., 2012; Schwager et al., 2017). There are ten Hox genes in all arthropods (Schwager et al., 2017). However, in some groups, like spiders, often two paralogs exist, as a result of a whole genome duplication (Schwager et al., 2017).

The expression of these genes has previously been studied in true spiders, among them the common house spider Parasteatoda tepidariorum and the Central American wandering spider Cupiennius salei (e.g. Damen, 1998; Khadjeh et al., 2012; Schwager et al. (2007, (2017))), but although the genes have been identified in A. geniculata (Janssen, unpublished data), and relationship between the genes in A. geniculata and P. tepidariorum has been determined with a phylogenetic analysis (Figure 1), and two of these genes, Antp-2 and AbdA-2 have been cloned (Janssen, unpublished data), there are no expression patterns of any tarantula Hox gene thus far.

Presence and relationship between the genes in A. geniculata and P. tepidariorum has been determined with a genetic analysis (Figure 1), and two of these genes, Antp-2 and AbdA-2 have been cloned, but there are no expression patterns of any tarantula Hox gene thus far.

This study focuses on the comparison of the four most posterior Hox genes, each present in the form of two paralogs, Antennapedia-1 (Ag-Antp-1) and Antennapedia-2 (Ag-Antp-2), Ultrabithorax-1 (Ag-Ubx-1) and Ultrabithorax-2 (Ag-Ubx-2), AbdominalA-1 (Ag-AbdA-1) and AbdominalA-2 (Ag-AbdA-2), AbdominalB-1 (Ag-AbdB-1) and AbdominalB-2 (Ag-AbdB-2). This is because these genes are involved in the formation of the opisthosomal structures book lungs (as well as tracheae in true spiders) and spinnerets (e.g. Schwager et al. 2017).

Figure 1: Phylogenetic tree of Hox genes, Antennapedia, Ultrabithorax, AbdominalA, and AbdominalB based on entire open reading frames (Janssen, unpublished data) to show the phylogenetic relationship between the genes in Parasteatoda tepidariorum and Acanthoscurria geniculata. It suggests that each posterior Hox gene in Pt possesses one ortholog in Ag.

Methods

Gene cloning The genes worked on from the start were Ag-Ubx-1, Ag-Ubx-2, Ag-AbdA-1, Ag-Antp-1 and Ag- AbdB-1. The others, Ag-AbdA-2, and Ag-Antp-2, were isolated before (Janssen, unpublished data). Primers (see Appendix A) for each gene were obtained from Macrogen and diluted to a 100mM solution. 1 µl of each primer (forward and backward primers) were added to separate tubes, along with 14.8 µl H2O, 2.5 µl 10x buffer solution, 2.5 µl of 2 mM dNTPs, 1.5 µl 25 mM MgCl2, 1 µl cDNA and 0.2 µl 5 U/µl Taq-polymerase. After the first PCR, a second PCR was done with the same ingredients, but with an inner pair of primers and 1 µl of the first PCR product instead of cDNA.

DNA ligation and bacterial transformation Two methods were used to ligate fragments into bacterial vectors. The first, used on Ag-AbdA- 1 and Ag-Ubx-2, was a topoisomerase ligation reaction. 3 µl of the PCR product, 1 µl H2O, 1 µl salt solution and 1 µl TOPO vector (ThermoFisher) were added to a tube and incubated at room temperature for 15 minutes. 2 µl of this mix was added to a tube of Escherichia coli on ice and incubated for 20 minutes.

The second method was ligation with a T4 ligase, this was done on Ag-AbdB-1 and Ag-Antp- 1. 2.5 µl of each gene from second PCR product was added to a tube each with 1 µl 5x buffer solution, 1 µl vector and 0.5 µl T4-DNA ligase. 2 µl were added to tubes with chemically competent cells on ice and incubated for 20 minutes.

Bacteria plus vectors were heat-shocked in a water bath at 42°C for 30 seconds, after which 250 µl of SOC-medium was added immediately after the tubes were briefly put into ice. This was done regardless of the previous ligation method. They were then incubated in a shaker at 37°C for one hour. The contents were added tospread on agar plates of with kanamycin treated with and 50µl X-gal in dimethylformamide and incubated overnight at 37°C. Kanamycin allows only bacteria that have been transformated (taken up a vector) to grow, because the kanamycin-resistance gene is located on the vector. X-gal allows for the distinguishing of self- ligated vectors from vectors with an insert (PCR fragment). Blue colonies are self-ligated, white colonies represent colonies with an insert; insertion of a PCR fragment destroys the beta- Gal gene located at the insertion site on the vector. This beta-Gal gene would be needed to metabolize the X-gal (the metabolic product is blue).

Colony PCR A master mix for each PCR was made, containing 64 µl H2O, 10 µl 10x buffer, 10 µl of 2mM dNTPs, 8 µl 25 mM MgCl2, 4 µl of 20 mM M13 forward primer, 4 µl of 20 mM M13 reverse primer and 0.5 µl 5 U/µl Taq-polymerase. This was then divided into eight PCR tubes (with 10 µl each of the master mix) per gene. Eight white colonies per plate were chosen and placed in a PCR tube each (and then dabbed on a control plate). A gel electrophoresis was run to determine whether the bacteria had picked up a fragment corresponding to the right size, and a maximum of two of those were chosen for subsequent sequencing. Chosen colonies were picked from the control plate with a pipette tip and placed into a tube (half-open lid for oxygen access) with 3 ml LB-medium + kanamycin. These were then incubated overnight in a shaker at 37℃.

Plasmid extraction To extract plasmids from the selected bacteria, a ThermoFisher Plasmid DNA miniprep kit was used following the instructions provided, with the following changes; the neutralisation solution was added after three minutes instead of five, and 80 µl elution solution was used instead of 100 µl. Concentration and purity was checked on a photometer.

Sequencing To determine that the inserted DNA fragments represent the desired fragments of Hox genes, the plasmids were sent to Macrogen for Sanger-sequencing. The resulting sequences were blasted to check their closest match, and the orientation of the fragments was confirmed to be positive (read forward) or negative (backward). This determined whether SP6 or T7 RNA polymerase should be used for subsequent RNA-probe synthesis.

Template PCR

A master mix was made, identical to the one done for colony PCR, but instead of adding colonies separately, 0.2 µl of the extracted plasmids were added directly to the master mix. This was then divided up into eight 0.2 ml tubes with 10 µl each to make a template PCR product. Plasmids cannot be used directly for RNA-probe synthesis because they are circular. One either has to digest (cut) them on the right side (depending on the RNA polymerase one needs to use to synthesise anti-sense RNA-probes, or a template PCR as described has to be performed with primers outside the RNA polymerase binding sites).

A ThermoFischer DNA purifying kit was used to clean PCR products according to the instructions from the manufacturer, with some changes. 40 µl of elution solution was used (once) instead of 50 µl.

Anti-sense RNA-probe synthesis 3 µl purified DNA, 3 µl H2O, 1 µl transcription buffer, 1 µl RNase inhibitor, 1 µl DIG labelling mix and 1 µl SP6 or T7 RNA polymerase was used (depending on orientation of the inserts (given PCR product inside the vector)). SP6 was used for the synthesis of anti-sense probes of Ag-Ubx-2, Ag-AbdA-1 Ag-AbdB-1 and Ag-Antp-1, and T7 was used for Ag-Ubx-1. The reactions were incubated at 37℃ for three hours. Afterwards, a QIAGEN RNeasy kit was used following instructions provided by the manufacturer to purify the synthesized probes.

Whole mount in-situ hybridization Fixed and long-time stored (in methanol) A. geniculata embryos were treated in 2% hydrogen peroxide in methanol for 20 minutes at room temperature. The embryos were then washed in 50% MeOH in phosphate buffered saline with tween (PBS-T) for 15 minutes (each time the liquid was replaced without letting the embryos touch air), followed by a 15 minute wash in 25% MeOH in PBS-T. They were then washed three times in PBS-T for 15 minutes, and postfixed for 20 minutes in 5% formaldehyde in PBS-T. Then they were washed another three times in PBS-T for 10 minutes each.

Embryos were washed once in a 1:1 solution of PBS-T and HYB-B (see Appendix B). This liquid was then replaced with pure HYB-B and the embryos were incubated for 15 minutes at 65℃. The HYB-B was then replaced with prewarmed HYB-A (see Appendix B), and the embryos were incubated at 65℃ for three hours.

The probes were then heated to 85℃ for two minutes, then placed on ice for two minutes and heated up to 65℃. The HYB-A was removed from the embryos and a fresh volume of 50 µl prewarmed HYB-A was added, along with 2.5 µl of the RNA-probes (one probe per tube). The embryos were then incubated overnight at 65℃.

The embryos were washed twice with 500 µl prewarmed HYB-B and incubated for 15 minutes at 65℃. The liquid was replaced with a 1:1 mixture of HYB-B and 2XSSC (see Appendix B) and incubated for 15 minutes at 65℃. Afterwards, they were washed twice with pure 2XSSC for 15 minutes each, at 65℃. Then another wash with PBS-T was done three times, ten minutes each at room temperature, and the embryos were incubated in blocking buffer for two hours at

room temperature. Blocking buffer is PBS-T that contains 2% sheep serum and 10mg/ml bovine serum albumine. This assures the blocking of nonspecific antibody-binding sites in the embryos.

A 1:2000 dilution of AP conjugated DIG-antibody with blocking buffer was prepared and the embryos were incubated with this mixture for two hours. They were then washed four times in PBS-T for 10 minutes each at room temperature, followed by an overnight incubation in PBS- T at 4℃.

The embryos were washed three times with PBS-T for ten minutes each, and then washed with alkaline-phosphate staining buffer (AP buffer, see Appendix B) three times for five minutes each. The embryos were then incubated with BM purple (Roche) staining solution for two hours in the dark. They were checked from time to time to follow the staining process. When appropriate the staining was stopped by washing the embryos four times for five minutes each with a less alkaline AP buffer (pH 7.4) (STOP solution); at this pH the AP does not work anymore. The embryos were counter-stained with CYBR Green in STOP-solution for 30 minutes in the dark. The unbound CYBR Green was then removed by washing three times in STOP solution, ten minutes each. 50 µl 37% FA was added to stained embryos in 1 ml STOP solution and stored at 4°C.

Data documentation The embryos were photographed under a light microscope. The CYBR Green signal was detected using a UV-lamp and the GFP-2 light filter.

Results

Antennapedia-1 There is a signal in the segment addition zone (SAZ) at stage 6 (using the staging system by Pechmann (2020)) (Figure 2a). At stage 8, the signal remains in the SAZ and extends to O1 in the form of stripes (Figure 2b). The signal remains in all opisthosomal segments at stage 10. (Figure 2c). At stage 11, the expression remains, although weaker, in all opisthosomal segments and is stronger in O1 and O2. Signal is also strong in the limb buds of segments O2 and O3 and spinneret buds of O4 and O5 (Figure 2g,h). At stage 12, the signal is limited to O1 and the anterior half of O2 as well as its limb bud (Figure 2i).

Antennapedia-2 Earliest expression discovered in stage 6 shows weak expression in O1 (Appendix C) Expression is present in the last walking leg, L4, and there is a weak expression in O1 and O2 at stage 11 (Figure 3a). At stage 12, the expression in L4 is larger, and the expression in O1 and O2 is limited to the dorsal areas (Figure 3b,c).

Ultrabithorax-1

There is a signal in the posterior part of the segment addition zone at stage 8 (Figure 4a). The signal is present in the opisthosomal segments posterior to O1 including in the limb and spinneret buds in segments O2-O5 (Figure 4b). At stage 12, the expression remains in the same segments, but the expression is only ventral. There is also still expression in the limb and spinneret buds, but the expression in the limb buds at O2 and O3 are stronger. The expression in the spinneret buds is also not present in the entire buds, but only the ventral part (Figure 4c). More images in Appendix D.

Ultrabithorax-2 Like Ag-Ubx-1, expression is solely in the opisthosomal area in the available stages. In stage 11, there is expression in the posterior part of O2 including the posterior part of its bud, with expression strong in O3 and O4, and weak expression in the posterior spinneret buds (Figure 5a,d). At stage 12, the signal is stronger and covers a larger area, but the expression remains strongest in O3 and O4 (Figure 5b,e). Later in stage 12, the strong signals in O3 and O4 remain as the posterior opisthosomal segments show weak expression (Figure 5c,f). More images in Appendix E.

AbdominalA-1 At stage 10, there is expression in the posterior part of O4, as well as O5 and O6, with a stronger expression in O5 (Figure 6a,c). At stage 11 (Figure 6b,d), the expression now covers the opisthosomal area from O4 to O8, including expression in the anterior and posterior spinneret buds, where it is stronger than the rest of segments O4 and O5.

AbdominalA-2 At stage 11, the signal in segments O5 to SAZ are strongest in the posterior ends, and there is a dorsal signal in O4. Signal is stronger between O5 and segment addition zone, with a signal in the dorsal area of O4 (Figure 7a). The expression is more homogenous in the segments by stage 12, but the expression in O5 is more dorsal than in the previous stage.

AbdominalB-1 At stage 9, there is a weak expression in the posterior part of O5 (Figure 9a,d). When the embryo reaches stage 10, the expression in O5 has expanded but remains in the posterior half of the segment, and there is expression in the posterior part of O6 and anterior part of O7 (Figure 9b,e). There is also a strong signal in the spinneret buds which gets stronger in stage 12. Additionally by stage 12, the expression has reached all segments posterior to O5 and the expression is equally strong in all segments (Figure 9c,f).

AbdominalB-2 Stage 8 of embryonic development shows a weak expression in the segment addition zone (Figure 9a.d). At stage 11 the expression has increased and has its border in the posterior of O5 as well as in the limb bud of O5 (Figure 9b,e). Later on in stage 11, the expression increases in strength but does not change location (Figure 9c,f).

Figure 2: Posterior side to the left in all pictures. a. Ventral view. Signal in segment addition zone at stage 6 (Pechmann, 2020) b. Ventral view. Signal remains in SAZ and extends to O1 in stripes at stage 8. c. Lateral view, ventral up. Signal strong in all opisthosomal segments at stage 10. d-f. CYBR green light photos of embryos a-c. g. Ventral view. At stage 11, the expression remains in all opisthosomal segments, but has become weaker. The expression is stronger in O1 and O2, including the limb bud of O2. Strong expression in the buds in O3-5 as well. h. Posterior view of embryo g. Signal is strong in the buds of segments O2-O5. i. Lateral view, ventral up. At

stage 12, the signal is limited to O1 and the posterior half of O2, including the limb bud. j-l. CYBR green light photos of embryos g-i.

Figure 3: All pictures of embryos with the posterior side facing left. a. Lateral view, ventral up. Strong signal in appendage of L4, as well as a weak signal in O1 and O2 at stage 11. b. Lateral view, ventral down. Expression in stage 12 is located in the same areas as stage 11, but the expression in the L4 appendage has grown larger. Several rings of expression. c. Dorsal view of embryo b. Expression in O2 is dorsal. d-f. Embryos a-c photographed in CYBR Green for identification of embryonic segments.

Figure 4: All pictures of embryos with the posterior side facing left. a. Ventral view. Strong signal in the posterior part of the segment addition zone at stage 8. b. Ventral view. Signal is present in all opisthosomal segments posterior to O1, including buds in O2-O5 at stage 11. Dots in L1-L3 are a stage-specific artefact and are not actual expressions of the gene. c. Lateral view, ventral down. Expression strong in ventral ectoderm. Expression in buds O2-O5 with stronger expression in O2 and O3. d-f. Pictures in CYBR Green light to easily identify segments.

Figure 5: All pictures taken from below with posterior end facing left. a. Stage 11 of embryonic development. Strong signal in segments O3-O4. with a slight extension to O2. Some expression in the spinneret buds of O5. b. Signal is at stage 12 still strongest in segments O3 and O4, but the expression is stronger posterior to that compared with stage 11. c. Signal is strongest in anterior parts of O3 and O4 at stage 12. d-f. Embryos photographed in CYBR Green for identification of embryonic segments.

Figure 6: All pictures with posterior end facing left. a. Ventral view. At stage 10, there is expression in O5 and O6 and the posterior part of O4. The expression in O5 is stronger than in both O6 and O4. b. Posterior view. Expression now covers opisthosomal area from O4 to O8 of the embryo at stage 11, with expression in both the anterior and posterior spinneret buds. c-d. Embryos photographed in CYBR Green for identification of embryonic segments.

Figure 7: All pictures with posterior view. a. Stage 11 of embryonic development. The signal in segments O5 to SAZ are strongest in the posterior ends, and there is a dorsal signal in O4. Signal is stronger between O5 and segment addition zone, with a signal in the dorsal area of O4. b. At stage 12 the expressions are in the same areas, but no longer predominantly in the posterior ends. The signal in O5 is however more dorsal than in stage 11. c- d. Embryos photographed in CYBR Green for identification of embryonic segments.

Figure 8: All pictures taken from below with posterior end facing left. a. Stage 9 of development shows expression in posterior part of O5. b. Expression in O5 is larger but still posterior by stage 10, expression strong in posterior spinneret buds. Expression in O6 stronger in the posterior area, and in O7 stronger in the anterior area. c. At stage 12, the expression is present in all segments posterior to O5, and expression in the posterior spinnerets are stronger. d-f. Embryos photographed in CYBR Green for identification of embryonic segments.

Figure 9: a. Ventral view of embryo at stage 8 in development. Weak expression in posterior area of segment addition zone. b. Posterior view. Expression in posterior part and in spinneret buds of O5 and in all segments posterior to O5 at stage 11. c. Lateral view of a late stage 11 embryo, ventral up. Expression increases in strength but remains in the same areas.

Discussion

This study showed clear expression patterns similar to those of P. tepidariorum and C. salei, and differences in morphology seem to be clearly related to the differences in Hox gene expression. Where there are no obvious differences in expression, there would need to be a difference somewhere down the gene cascades. These differences can not be identified in this study as Hox gene expression is the only thing looked at. To identify such differences, one would need to study the cascades in detail in compared species.

Antennapedia In available embryos, the earliest expression of Antp-1 found in A. geniculata was in the segment addition zone (SAZ) in stage 6 (Figure 2a). This is one stage earlier than the expression was initially found in P. tepidariorum. The expression in all three spiders remains in every segment in the opisthosoma, but in P. tepidariorum and C. salei, the expression also reaches the posterior half of L4. This is not seen in the available data for A. geniculata. It could be the case that there is not enough data to definitively say, but P. tepidariorum shows a signal in L4 as early as stage 9, which is clearly not the case in A. geniculata. The expression of P. tepidariorum appears in the form of stripes from the SAZ, and new stripes appear until segmentation is complete. This can be seen in A. tepidariorum as well. By stage 9, the expression in P. tepidariorum becomes weaker, but there is a strong expression in the ventral neural ectoderm and expression in the SAZ. While the expression becomes weaker in the available later stages of A. geniculata, there is no significant expression in the neural ectoderm, and the O2-O5 buds keep a strong signal that they do not in P. tepidariorum. A common trait between all three spiders is the expression of the gene in stages 11 and 12. Around this point in development, all three spiders show a strong signal in O1 and the anterior half of O2. In A. geniculata, this can be seen at stage 12, whereas P. tepidariorum and C. salei show this at stage 11. A difference between A. geniculata and the other two spiders is that in A. geniculata, the limb bud expression at stage 12 remains strong while it is completely gone at stage 11 of P. tepidariorum and C. salei.

Antp-2 in A. geniculata is expressed strongly in the appendage of L4 as well as O1 and O2. In P. tepidariorum, expression begins in segments O1 and O2 at stage 8. The definitive start of expression in A. geniculata is unclear, but the earliest available is in stage 6 with a weak expression in O1. In both P. tepidariorum and A. geniculata, the expression extends into the L4 appendage in the form of rings. From L4, the expression border in P. tepidariorum reaches O3, but in A. geniculata the expression only extends to the border of O2.

Earlier studies of C. salei indicated that while the Antp paralogs form legs in insects, the same function in spiders is unlikely because legs do not form where the genes are active in spiders

(Khadjeh et al., 2012), namely the opisthosomal segments posterior to the walking legs. This is strengthened by the similar results found in A. geniculata. It’s interesting that Antp-2 is expressed in L4 despite those genes being repressors, this could be explained in a few ways. It’s possible that it does not repress leg growth if combined with the other leg forming Hox genes, Sex comb reduced that are active in L4. Another, more likely reason, is the temporal factor. Perhaps repression simply does not occur if Antp-2 reaches L4 after the forming of the leg has already started.

Ultrabithorax A. geniculata expresses Ubx-1 initially in the posterior part of the segment addition zone, with the earliest stage available being stage 8. The same is true of P. tepidariorum, but P. tepidariorum, unlike A. geniculata, also shows a weak expression in a band anterior to the SAZ. This anterior band is present in multiple stages of embryonic development in P. tepidariorum, but is not present in any available stage in A. geniculata (see Appendix D). Interestingly, C. salei seems to lack this band as well. In all available stages, A. geniculata has a strong expression of Ubx-1 in the buds of each segment. once the expression reaches O2 (where the expression border is anterior in all three spiders), even as the signal weakens and is expressed in the ectoderm in stage 12, the expression in the buds remains strong. The expression in the buds is strong in C. salei as well as P. tepidariorum, but as the signal weakens in P. tepidariorum and finds the ventral neuroectoderm, the buds do not show a strong signal to the extent it is shown in A. geniculata.

Ubx-2 in A. geniculata is shown initially (earliest available stage is stage 11) in segments O2 (although only posterior half) to O4 in available pictures, with a weak expression in the posterior spinneret buds. By stage 12, one stage later, the expression remains strongest in O3 and O4 but is expressed posterior to that as well. All three spiders have the expression border in the posterior half of O2. This is not surprising as all three spiders have book lungs in O2. However, where A. geniculata has book lungs in O3, there are tracheae in true spiders instead.

Studies on Ubx in Drosophila melanogaster shows that the gene has a repressing function in the case of halteres, where Ubx controls the size (de Navas et al., 2006). This could imply that Ubx has a similar role in the development of book lungs and tracheae. There is no obvious difference in the expression patterns of Ubx-2 in either spider, however Ubx-1 in A. geniculata remains in O2 and O3 longer than in P. tepidariorum. It is likely then that when the Ubx expression in P. tepidariorum is reduced, it is possible for it to develop tracheae instead of book lungs.

AbdominalA and Abdominal B Because the posterior spinnerets are reduced in Mygalomorphae (and not in Araneomorphae), we would expect to see a difference in expression in the Abdominal genes if they have a role in spinneret development.

AbdA-1 in P. tepidariorum is expressed further than in A. geniculata. In A. geniculata, the expression border is in segment O4 (the expression border is in O2 in P. tepidariorum). This is

curious as O4 is the location of the anterior spinnerets that remain in P. tepidariorum but are reduced in A. geniculata. The expression is also not weak, so a possible explanation is that AbdA-1 and AbdB-1 are needed together in some capacity to form spinnerets. This makes sense as there is expression of AbdB-1 in P. tepidariorum in O4 (albeit only in the dorsal area), but that may well be enough to give this segment its identity, whereas A. geniculata has a very strong expression of Ag-AbdB-1 in O5 (potentially explaining the enlarged spinnerets compared to those in Araneomorphae),but does not at all reach O4. In C. salei, the anterior expression border of Abd is found in the posterior half of O3, including the posterior half of the O3 limb bud. Which paralog is unclear, so incorporating the Hox gene into the tree (Figure 1) would be necessary to determine this.

AbdA-2 in A. geniculata does reach O4, but is only in the dorsal area. In P. tepidariorum the expression is equally strong in O4 and posterior to that, so that too suggests that the limit of AbdB-1 in A. geniculata determines the fate of the spinnerets.

AbdB-2 has a similar expression pattern in A. geniculata and P. tepidariorum with an initial weak expression in the SAZ at stage 6 (Appendix F). However, like in Ubx-1, P. tepidariorum shows a weak band of expression further up in the opisthosoma that is not there in A. geniculata.The expression border in A. geniculata is in the posterior part of O5, where there is also expression in the O5 spinneret buds. P. tepidariorum eventually shows expression in the O2 limb buds, that does not appear in A. geniculata. It does appear in C. salei where, aside from the expression in the limb bud of O2, the expression extends from the anterior half of O4 back to the posterior end of the embryo. AbdB-2 also appears in the ventral neuroectoderm of P. tepidariorum up to O3, but in all available stages of A. geniculata, the expression does not extend anterior to O5. This may be evidence that AbdB-2 expression too is important for the development of anterior spinneret buds.

Potential improvements Because the development stage is an important factor, having embryos in each stage to note the expression timeline is important. There were not enough embryos to see every stage in development, so the precise start of expression and end is unclear for all genes.

The use of RNAi to knock out certain Hox gene expressions in live embryos is a logical next move to see how morphology changes with inactive genes and would show direct evidence of the purpose of these genes in A. geniculata.

References:

Barnett AA, Thomas RH. 2013. Posterior Hox gene reduction in an arthropod: Ultrabithorax and Abdominal-B are expressed in a single segment in the mite Archegozetes longisetosus. EvoDevo 4, 23. https://doi.org/10.1186/2041-9139-4-23

Damen WGM, Hausdorf M, Seyfarth E et al. 1998. A Conserved Mode of Head Segmentation in Arthropods Revealed by the Expression Pattern of Hox Genes in a Spider. PNAS 95: 10665- 70. https://doi.org/10.1073/pnas.95.18.10665. de Navas LF, Garaulet DL, Sánchez-Herrero E. 2006. The Ultrabithorax Hox gene of Drosophila controls haltere size by regulating the Dpp pathway. Development 133: 4495-4506 https://doi.org/10.1242/dev.02609

Hughes CL, Kaufman TC. Hox genes and the evolution of the arthropod body plan. EvoDevo 4:459-99.

Khadjeh S, Turetzek N, Pechmann M et al. 2012. Divergent role of the Hox gene Antennapedia in spiders is responsible for the convergent evolution of abdominal limb repression. PNAS 109: 4921-4926. https://doi.org/10.1073/pnas.1116421109

Küntzel N, Dunlop JA, Scholtz G. 2019. Morphology and evolution of spider book lungs (Araneae).Arthropod Systematics & Phylogeny 77: 267-284. https://doi.org/10.26049/ASP77- 2-2019-05

Mariano-Martins P, Lo-Man-Hung N, Torres TT. 2020. Evolution of Spiders and Silk Spinning: Mini Review of the Morphology, Evolution, and Development of Spiders’ Spinnerets. Front. Ecol. Evol 8: 109. https://doi.org/10.3389/fevo.2020.00109

Pace RM, Grbić M, Nagy LM. 2016. Composition and genomic organization of arthropod Hox clusters. EvoDevo 7, 11. https://doi.org/10.1186/s13227-016-0048-4

Pechmann M, Prpic NM. 2009. Appendage patterning in the South American bird spider Acanthoscurria geniculata (Araneae: Mygalomorphae). Development Genes and Evolution 219: 189-198. https://doi.org/10.1007/s00427-009-0279-7

Pechmann M. 2020. Embryonic development and secondary axis induction in the Brazilian white knee tarantula Acanthoscurria geniculata, C. L. Koch, 1841 (Araneae; Mygalomorphae; Theraphosidae). Development Genes and Evolution 230: 75-94. https://doi.org/10.1007/s00427-020-00653-w

Schwager EE, Schoppmeier M, Pechmann M, Damen WGM. 2007. Duplicated Hox genes in the spider Cupiennius salei. Front Zool 4, 10. https://doi.org/10.1186/1742-9994-4-10

Schwager EE, Sharma PP, Clarke T et al. 2017. The house spider genome reveals an ancient whole-genome duplication during arachnid evolution. BMC Biol 15, 62. https://doi.org/10.1186/s12915-017-0399-x

Sharma PP, Schwager EE, Extavour CG et al. 2012. Hox gene expression in the harvestman Phalangium opilio reveals divergent patterning of the chelicerate opisthosoma. Evol Dev 14: 450-463. https://doi.org/10.1111/j.1525-142X.2012.00565.x

Wheeler WC, Coddington JA, Crowley LM et al. 2016. The spider tree of life: Phylogeny of Araneae based on target-gene analyses from an extensive taxon sampling. Cladistics 33: 574- 616

Appendix

Gene First PCR Second PCR

Ag-Ubx-1 Forward primer (Fw): Fw: 5’ 5’ - - CGAACAGAGCGGTTTCT CCACAGATCAGCCGTAT ACA-3’ AGA-3’

Backward primer (Bw): Bw: 5’- 5’- TCCATACGTTTTTGCAG CGACTGTTGTTGAGACT CCG-3’ TCT-3’

Ag-Ubx-2 Fw: Fw: 5’- 5’- CGAGCAACCCTACCGAT CAAGGACTGCTCGTACC TTC-3’ CC-3’

Bw: Bw: 5’- 5’- TTACGTTTTGGCATCGA ATAGCGGTGGTCTTGGT CCG-3’ CGT-3’

Ag-AbdA-1 Fw: Fw: 5’- 5’- TTATGGACACGTTTTTG ATGCCGTATCATCGCTG GGG-3’ TCA-3’

Bw: Bw: 5’- 5’- TTATGGCAAACTCGCTT GTCTGAGCTGGGTATTA CGT-3’ CAG-3’

Ag-AbdA-2 Fw: Fw: 5’- 5’- TGGCTACTCCTTCAACT CCTTCTCAGTTCTTCCCT

TGC-3’ CA-3’

Bw: Bw: 5’- 5’- TTTAATGAGCAGATCCC GATGAAATGGCTGATCT GAG-3’ GGA-3’

Ag-AbdB-1 Fw: Fw: 5’- 5’- GAACGGAAGTTTGTATG TCGAAGTCGTTCAGCCA AGG-3’ TCT-3’

Bw: Bw: 5’- 5’- TTACTTCACTGCTGTTG TCTGTCCGTTAGTAGCA TCG-3’ GCA-3’

Ag-Antp-1 Fw: Fw: 5’- 5’- TCGAAATGGAGGTGGA AACAACCACAGCAGCA GAAC-3’ GCAA-3’

Bw: Bw: 5’- 5’- TATGATGAGCGAGTTCA CCCATTATGAGACCCGA GGT-3’ TTC-3’

Ag-Antp-2 Fw: Fw: 5’- 5’- ACTTACGGAGTAGCGGT AGTCCATACGCAGCCCA GAA-3’ ATG-3’

Bw: Bw: 5’- 5’- GTTTGGCTTTGTTCTCCT TATTTTGATCTGCCTCT TC-3’ CCG-3’ A: Primer sequences used for Ag-Ultrabithorax-1 (Ag-Ubx-1), Ag-Ubx-2, Ag-AbdominalA-1 (Ag-AbdA-1), Ag-AbdB-1, Ag-AbdA-2, Ag-Antennapedia-1 (Ag-Antp-1) and Ag-Antp-2.

In-situ hybridisation solutions Ingredients and steps

•••10x PBS 1.37 M NaCl

27 mM KCl

100 mM Na2HPO4

Adjust pH to 7.4

Autoclave

PBS-T 1 x PBS

0.1 % Tween-20

HYB-B 50% formamide

25% 20x SSC, pH 7.0

0.1% Tween-20

Adjust pH to 6.5

HYB-A 1x HYB-B

0.05 mg/ml heparin

5% Dextran Sulfate Salt (Sigma) (2.5g in 50 ml)

0.01 mg/ml yeast RNA (or 250 µl 20mg/ml tRNA)

0.4 mg/ml sonicated salmon sperm DNA

Boil for 10 min and cool on ice for 3 min

Store @ -20°C

Blocking buffer 1x PBS-T

1% BSA

2% Sheep serum

AP staining buffer 100 mM Tris pH 9.5

150 mM NaCl

10 mM MgCl2

0.1% Tween-20

STOP buffer Same as AP staining buffer, pH 7.4

B: All components used in in-situ hybridisation and instructions.

C: Earliest available expression of Ag-Antp-2, stage 6. Weak expression in O1.

D: Additional images of Ag-Ubx-1 in no particular order.

E: Additional images of Ag-Ubx-2 in no particular order.

F: Ag-AbdB-2 expression at stage 6.