<figure><img src="image_1.png" /></figure>

HAL
open science

# γ-TuRCs and Augmin are required for the development of highly branched dendritic arbors in Drosophila

Amrita Mukherjee, Yaiza Andrés Jeske, Isabelle Becam, Anaelle Taïeb, Paul Brooks, Joanna Aouad, Clementine Monguillon, Paul T Conduit

<figure><img src="image_2.png" /></figure>

## To cite this version:

Amrita Mukherjee, Yaiza Andrés Jeske, Isabelle Becam, Anaelle Taïeb, Paul Brooks, et al.. γ-TuRCs and Augmin are required for the development of highly branched dendritic arbors in Drosophila. Journal of Cell Science, In press. hal-04534582

HAL Id: hal-04534582

https://hal.science/hal-04534582

Submitted on 5 Apr 2024

HAL is a multi-disciplinary open access archive for the deposit and dissemination of scientific research documents, whether they are published or not. The documents may come from teaching and research institutions in France or abroad, or from public or private research centers.

L'archive ouverte pluridisciplinaire HAL, est destinée au dépôt et à la diffusion de documents scientifiques de niveau recherche, publiés ou non, émanant des établissements d'enseignement et de recherche français ou étrangers, des laboratoires publics ou privés.

<figure><img src="image_3.png" /></figure>

Distributed under a Creative Commons Attribution 4.0 International License
<figure><img src="image_1.png" /></figure>

HAL
open science

# γ-TuRCs and Augmin are required for the development of highly branched dendritic arbors in Drosophila

Amrita Mukherjee, Yaiza Andrés Jeske, Isabelle Becam, Anaelle Taïeb, Paul Brooks, Joanna Aouad, Clementine Monguillon, Paul T Conduit

<figure><img src="image_2.png" /></figure>

## To cite this version:

Amrita Mukherjee, Yaiza Andrés Jeske, Isabelle Becam, Anaelle Taïeb, Paul Brooks, et al.. γ-TuRCs and Augmin are required for the development of highly branched dendritic arbors in Drosophila. Journal of Cell Science, 2024. hal-04534582

HAL Id: hal-04534582

https://hal.science/hal-04534582v1

Submitted on 5 Apr 2024

HAL is a multi-disciplinary open access archive for the deposit and dissemination of scientific research documents, whether they are published or not. The documents may come from teaching and research institutions in France or abroad, or from public or private research centers.

L'archive ouverte pluridisciplinaire HAL, est destinée au dépôt et à la diffusion de documents scientifiques de niveau recherche, publiés ou non, émanant des établissements d'enseignement et de recherche français ou étrangers, des laboratoires publics ou privés.

<figure><img src="image_3.png" /></figure>

Distributed under a Creative Commons Attribution 4.0 International License
1

γ-TuRCs and Augmin are required for the development of highly

2

branched dendritic arbors in *Drosophila*

3

4

Amrita Mukherjee*¹,², Yaiza Andrés Jeske*¹, Isabelle Becam³, Anaelle Taïeb³, Paul

5

Brooks¹, Joanna Aouad³, Clementine Monguillon³, and Paul T. Conduit³,⁴

6

7

¹Department of Zoology, University of Cambridge, Downing Street, Cambridge, CB2

8

3EJ

9

²MRC Toxicology Unit, Gleeson Building, Tennis Court Road, Cambridge CB2 1QR,

10

UK

11

³Université Paris Cité, CNRS, Institut Jacques Monod, F-75013, Paris, France.

12

*These authors contributed equally to the work

13

14

⁴Corresponding author

15

Paul T. Conduit

16

Institut Jacques Monod

17

CNRS - Université Paris Cité

18

15 rue Hélène Brion

19

75013 Paris

20

France

21

0033 (1) 57 27 80 95

22

paul.conduit@ijm.fr

23

ORCID: 0000-0002-7822-1191

24
1

γ-TuRCs and Augmin are required for the development of highly

2

branched dendritic arbors in *Drosophila*

3

4

Amrita Mukherjee*¹,², Yaiza Andrés Jeske*¹, Isabelle Becam³, Anaelle Taïeb³, Paul

5

Brooks¹, Joanna Aouad³, Clementine Monguillon³, and Paul T. Conduit³,⁴

6

7

¹Department of Zoology, University of Cambridge, Downing Street, Cambridge, CB2

8

3EJ

9

²MRC Toxicology Unit, Gleeson Building, Tennis Court Road, Cambridge CB2 1QR,

10

UK

11

³Université Paris Cité, CNRS, Institut Jacques Monod, F-75013, Paris, France.

12

*These authors contributed equally to the work

13

14

⁴Corresponding author

15

Paul T. Conduit

16

Institut Jacques Monod

17

CNRS - Université Paris Cité

18

15 rue Hélène Brion

19

75013 Paris

20

France

21

0033 (1) 57 27 80 95

22

paul.conduit@ijm.fr

23

ORCID: 0000-0002-7822-1191

24
25

**Abstract**

26

27

28

29

30

31

32

33

34

35

36

37

38

39

40

41

Microtubules are nucleated by γ-tubulin ring complexes (γ-TuRCs) and are essential for neuronal development. Nevertheless, γ-TuRC depletion has been reported to perturb only higher-order branching in elaborated *Drosophila* larval class IV dendritic arborisation (da) neurons. This relatively mild phenotype was attributed to defects in microtubule nucleation from Golgi outposts, yet most Golgi outposts lack associated γ-TuRCs. By analysing dendritic arbor regrowth in pupae, we show that γ-TuRCs are also required for the growth and branching of primary and secondary dendrites, as well as for higher-order branching. Moreover, we identify the Augmin complex, which recruits γ-TuRCs to the sides of pre-existing microtubules, as being required predominantly for higher-order branching, likely by increasing the frequency of microtubule growth in terminal dendrites. Consistently, we find that Augmin is expressed less strongly and is largely dispensable in larval class I da neurons, which exhibit few higher-order dendrites. Thus, γ-TuRCs are essential for various aspects of complex dendritic arbor development, and they appear to function in higher-order branching via the Augmin pathway, which promotes the elaboration of dendritic arbors to help define neuronal morphology.
25

Abstract

26

27

28

29

30

31

32

33

34

35

36

37

38

39

40

41

Microtubules are nucleated by γ-tubulin ring complexes (γ-TuRCs) and are essential for neuronal development. Nevertheless, γ-TuRC depletion has been reported to perturb only higher-order branching in elaborated *Drosophila* larval class IV dendritic arborisation (da) neurons. This relatively mild phenotype was attributed to defects in microtubule nucleation from Golgi outposts, yet most Golgi outposts lack associated γ-TuRCs. By analysing dendritic arbor regrowth in pupae, we show that γ-TuRCs are also required for the growth and branching of primary and secondary dendrites, as well as for higher-order branching. Moreover, we identify the Augmin complex, which recruits γ-TuRCs to the sides of pre-existing microtubules, as being required predominantly for higher-order branching, likely by increasing the frequency of microtubule growth in terminal dendrites. Consistently, we find that Augmin is expressed less strongly and is largely dispensable in larval class I da neurons, which exhibit few higher-order dendrites. Thus, γ-TuRCs are essential for various aspects of complex dendritic arbor development, and they appear to function in higher-order branching via the Augmin pathway, which promotes the elaboration of dendritic arbors to help define neuronal morphology.
42 **Introduction**

43 Microtubules are essential cytoskeletal components necessary for neuronal growth
44 and function (Kapitein and Hoogenraad, 2015; Rolls et al., 2021). These polarised
45 filaments run throughout axons and dendrites performing at least two important
46 functions. They provide tracks for molecular motors to traffic cargo between the soma
47 and neurite terminals, and their outward growth supports and facilitates neurite growth
48 and branching. Within axons, most microtubules have their plus ends pointing away
49 from the soma, referred to as plus-end-out microtubules, while dendrites contain either
50 mixed or predominantly minus-end-out microtubules (Rolls and Jegla, 2015). This
51 difference in microtubule polarity is important for the correct polarised trafficking of
52 cargos into axons and dendrites and thus for axon and dendrite identity (Rolls, 2011;
53 Tas et al., 2017; Thyagarajan et al., 2022).
54

54

55 The *de novo* formation of microtubules is kinetically unfavourable at physiological
56 concentrations of tubulin dimers. Thus, microtubule nucleation is promoted by
57 microtubule-associated proteins (MAPs) or protein complexes that locally concentrate
58 tubulin dimers (Tovey and Conduit, 2018). The best-characterised nucleator is the
59 multi-protein γ-tubulin ring complex (γ-TuRC), which templates the lateral assembly
60 of tubulin dimers to form a 13-protofilament microtubule that extends with its plus end
61 growing away from the γ-TuRC (Kollman et al., 2011). γ-TuRC activity is normally low
62 in the cytosol but increases on recruitment to specific sites called microtubule
63 organising centres (MTOCs) (Sanchez and Feldman, 2016; Tillery et al., 2018).
64 Centrosomes are the best-known and best-characterised MTOC and are very active
65 during mitosis (Conduit et al., 2015), but are often inactivated in neurons, particularly
66 after early developmental stages (Nguyen et al., 2011; Stiess et al., 2010). Thus, non-
67 centrosomal MTOCs play an important role in organising the neuronal microtubule
68 cytoskeleton (Lüders, 2020). So far, the surface of Golgi structures (Fu et al., 2019;
69 Ori-McKenney et al., 2012; Valenzuela et al., 2020; Yalgin et al., 2015; Yang and
70 Wildonger, 2020; Zhou et al., 2014)), endosomes (Weiner et al., 2020) and
71 centrosomal material in the distal tip of ciliated neurons (Garbrecht et al., 2021;
72 Harterink et al., 2018; Magescas et al., 2021) have been reported as non-centrosomal
42 **Introduction**

43 Microtubules are essential cytoskeletal components necessary for neuronal growth
44 and function (Kapitein and Hoogenraad, 2015; Rolls et al., 2021). These polarised
45 filaments run throughout axons and dendrites performing at least two important
46 functions. They provide tracks for molecular motors to traffic cargo between the soma
47 and neurite terminals, and their outward growth supports and facilitates neurite growth
48 and branching. Within axons, most microtubules have their plus ends pointing away
49 from the soma, referred to as plus-end-out microtubules, while dendrites contain either
50 mixed or predominantly minus-end-out microtubules (Rolls and Jegla, 2015). This
51 difference in microtubule polarity is important for the correct polarised trafficking of
52 cargos into axons and dendrites and thus for axon and dendrite identity (Rolls, 2011;
53 Tas et al., 2017; Thyagarajan et al., 2022).
54

54

55 The *de novo* formation of microtubules is kinetically unfavourable at physiological
56 concentrations of tubulin dimers. Thus, microtubule nucleation is promoted by
57 microtubule-associated proteins (MAPs) or protein complexes that locally concentrate
58 tubulin dimers (Tovey and Conduit, 2018). The best-characterised nucleator is the
59 multi-protein γ-tubulin ring complex (γ-TuRC), which templates the lateral assembly
60 of tubulin dimers to form a 13-protofilament microtubule that extends with its plus end
61 growing away from the γ-TuRC (Kollman et al., 2011). γ-TuRC activity is normally low
62 in the cytosol but increases on recruitment to specific sites called microtubule
63 organising centres (MTOCs) (Sanchez and Feldman, 2016; Tillery et al., 2018).
64 Centrosomes are the best-known and best-characterised MTOC and are very active
65 during mitosis (Conduit et al., 2015), but are often inactivated in neurons, particularly
66 after early developmental stages (Nguyen et al., 2011; Stiess et al., 2010). Thus, non-
67 centrosomal MTOCs play an important role in organising the neuronal microtubule
68 cytoskeleton (Lüders, 2020). So far, the surface of Golgi structures (Fu et al., 2019;
69 Ori-McKenney et al., 2012; Valenzuela et al., 2020; Yalgin et al., 2015; Yang and
70 Wildonger, 2020; Zhou et al., 2014)), endosomes (Weiner et al., 2020) and
71 centrosomal material in the distal tip of ciliated neurons (Garbrecht et al., 2021;
72 Harterink et al., 2018; Magescas et al., 2021) have been reported as non-centrosomal
MTOCs in *Drosophila* and *C. elegans* neurons. Microtubules are also nucleated from *en passant* boutons in cultured mammalian hippocampal neurons (Qu et al., 2019).

Another important mode of non-centrosomal microtubule nucleation that has been implicated in neuronal development is nucleation via the Augmin complex (known as the HAUS complex in mammals). Each Augmin complex recruits individual γ-TuRCs to the sides of pre-existing microtubules to nucleate an individual microtubule with the same polarity as the mother microtubule (Goshima et al., 2008; Kamasaki et al., 2013; Lawo et al., 2009; Liu et al., 2014; Petry et al., 2013; Song et al., 2018; Verma and Maresca, 2019). These sparsely distributed nucleation events can be parallel or branched depending on the cell type and the presence or absence of certain protein and protein modifications (Sánchez-Huertas and Lüders, 2015). Augmin subunits were first identified in a screen for factors required for spindle assembly in *Drosophila* S2 cells and were shown to be important for microtubule generation within the mitotic spindle (Goshima et al., 2007; Goshima et al., 2008). This was consistent with previous observations of microtubule nucleation from pre-existing microtubules within the cortical network of plant and fission yeast interphase cells (Janson et al., 2005; Murata et al., 2005; Wasteneys and Williamson, 1989), which was subsequently also observed in Xenopus egg extracts (Petry et al., 2013). The molecular architecture of Augmin has now been revealed by a combination of cryo-EM and computational predictions, providing molecular details on how these complexes simultaneously bind microtubules and γ-TuRCs (Gabel et al., 2022; Zupa et al., 2022). Direct observations of single Augmin-mediated nucleation events have also been made using TIRF microscopy in *Drosophila* S2 cells (Verma and Maresca, 2019). In mammalian neurons, Augmin depletion affects cortical neuron migration and polarisation in mice and microtubule density and polarity in the dendrites and axons, respectively, of cultured rat hippocampal neurons (Cunha-Ferreira et al., 2018; Sánchez-Huertas et al., 2016). Nevertheless, whether neurons have a general requirement for Augmin, or whether it is required in specific settings or for specific processes remains unclear.

*Drosophila* larval dendritic arborisation neurons (da neurons) are a well-established model system to study neuronal morphology and microtubule regulation *in vivo* (Jan
MTOCs in *Drosophila* and *C. elegans* neurons. Microtubules are also nucleated from *en passant* boutons in cultured mammalian hippocampal neurons (Qu et al., 2019).

Another important mode of non-centrosomal microtubule nucleation that has been implicated in neuronal development is nucleation via the Augmin complex (known as the HAUS complex in mammals). Each Augmin complex recruits individual γ-TuRCs to the sides of pre-existing microtubules to nucleate an individual microtubule with the same polarity as the mother microtubule (Goshima et al., 2008; Kamasaki et al., 2013; Lawo et al., 2009; Liu et al., 2014; Petry et al., 2013; Song et al., 2018; Verma and Maresca, 2019). These sparsely distributed nucleation events can be parallel or branched depending on the cell type and the presence or absence of certain protein and protein modifications (Sánchez-Huertas and Lüders, 2015). Augmin subunits were first identified in a screen for factors required for spindle assembly in *Drosophila* S2 cells and were shown to be important for microtubule generation within the mitotic spindle (Goshima et al., 2007; Goshima et al., 2008). This was consistent with previous observations of microtubule nucleation from pre-existing microtubules within the cortical network of plant and fission yeast interphase cells (Janson et al., 2005; Murata et al., 2005; Wasteneys and Williamson, 1989), which was subsequently also observed in Xenopus egg extracts (Petry et al., 2013). The molecular architecture of Augmin has now been revealed by a combination of cryo-EM and computational predictions, providing molecular details on how these complexes simultaneously bind microtubules and γ-TuRCs (Gabel et al., 2022; Zupa et al., 2022). Direct observations of single Augmin-mediated nucleation events have also been made using TIRF microscopy in *Drosophila* S2 cells (Verma and Maresca, 2019). In mammalian neurons, Augmin depletion affects cortical neuron migration and polarisation in mice and microtubule density and polarity in the dendrites and axons, respectively, of cultured rat hippocampal neurons (Cunha-Ferreira et al., 2018; Sánchez-Huertas et al., 2016). Nevertheless, whether neurons have a general requirement for Augmin, or whether it is required in specific settings or for specific processes remains unclear.

*Drosophila* larval dendritic arborisation neurons (da neurons) are a well-established model system to study neuronal morphology and microtubule regulation *in vivo* (Jan
and Jan, 2010). They form in embryos but continue to develop throughout the larval stages until pupation. They exhibit elaborate but stereotypical dendritic arbors that grow mainly in two dimensions in the larval body wall, making them amenable to morphology analysis and accessible to live imaging *in situ*. They are genetically tractable with well-characterised neuron-type-specific gal4 drivers able to express UAS-controlled fluorescent reporters and RNAi constructs. Da neurons are split into four classes (Grueber et al., 2002), with simple class I and elaborate class IV neurons most commonly studied. Class I neurons are proprioceptive and comprise a relatively long primary dendrite that branches repeatedly into multiple secondary dendrites that extend along the antero-posterior axis to sense stretch during larval crawling (Song et al., 2007; Vaadia et al., 2019). These secondary dendrites generate few higher-order dendritic branches. In contrast, class IV da neurons are nociceptive e.g. (Hu et al., 2017; Xiang et al., 2010; Zhong et al., 2010) and tile the body wall by branching extensively (Grueber et al., 2002). They comprise long primary and secondary dendrites that branch into many tertiary and terminal (higher-order) branches to cover their receptive field.

Within the dendrites of class I and class IV da neurons, it was proposed that γ-TuRCs associate with fragments of Golgi called Golgi outposts to mediate de-centralised microtubule nucleation events that regulate microtubule polarity and arbor morphology (Ori-McKenney et al., 2012; Yalgin et al., 2015). The main phenotype after γ-tubulin depletion from class IV neurons was a reduction in the number and length of higher-order branches and it was concluded that this was due to defects in γ-TuRC-mediated microtubule nucleation from Golgi outposts (Ori-McKenney et al., 2012). However, the conclusion that Golgi outposts help regulate microtubule polarity in da neurons was later challenged by findings showing that exogenously-expressed γ-tubulin-GFP localises to branch points independently of Golgi outposts and that microtubule polarity was unaffected by the re-positioning of Golgi outposts out of dendrites (Nguyen et al., 2014). Using endogenously regulated reporters of γ-TuRCs, it was then shown that γ-TuRCs localise to relatively few Golgi outposts that are present only within primary branch points (close to the soma) (Mukherjee et al., 2020). These γ-TuRC-positive Golgi outposts are not directly associated with terminal branches,
and Jan, 2010). They form in embryos but continue to develop throughout the larval stages until pupation. They exhibit elaborate but stereotypical dendritic arbors that grow mainly in two dimensions in the larval body wall, making them amenable to morphology analysis and accessible to live imaging *in situ*. They are genetically tractable with well-characterised neuron-type-specific gal4 drivers able to express UAS-controlled fluorescent reporters and RNAi constructs. Da neurons are split into four classes (Grueber et al., 2002), with simple class I and elaborate class IV neurons most commonly studied. Class I neurons are proprioceptive and comprise a relatively long primary dendrite that branches repeatedly into multiple secondary dendrites that extend along the antero-posterior axis to sense stretch during larval crawling (Song et al., 2007; Vaadia et al., 2019). These secondary dendrites generate few higher-order dendritic branches. In contrast, class IV da neurons are nociceptive e.g. (Hu et al., 2017; Xiang et al., 2010; Zhong et al., 2010) and tile the body wall by branching extensively (Grueber et al., 2002). They comprise long primary and secondary dendrites that branch into many tertiary and terminal (higher-order) branches to cover their receptive field.

Within the dendrites of class I and class IV da neurons, it was proposed that γ-TuRCs associate with fragments of Golgi called Golgi outposts to mediate de-centralised microtubule nucleation events that regulate microtubule polarity and arbor morphology (Ori-McKenney et al., 2012; Yalgin et al., 2015). The main phenotype after γ-tubulin depletion from class IV neurons was a reduction in the number and length of higher-order branches and it was concluded that this was due to defects in γ-TuRC-mediated microtubule nucleation from Golgi outposts (Ori-McKenney et al., 2012). However, the conclusion that Golgi outposts help regulate microtubule polarity in da neurons was later challenged by findings showing that exogenously-expressed γ-tubulin-GFP localises to branch points independently of Golgi outposts and that microtubule polarity was unaffected by the re-positioning of Golgi outposts out of dendrites (Nguyen et al., 2014). Using endogenously regulated reporters of γ-TuRCs, it was then shown that γ-TuRCs localise to relatively few Golgi outposts that are present only within primary branch points (close to the soma) (Mukherjee et al., 2020). These γ-TuRC-positive Golgi outposts are not directly associated with terminal branches,
137

suggesting that the loss of terminal branches after γ-tubulin depletion may represent
a failure in an as yet unidentified, Golgi outpost-independent, mode of microtubule
nucleation. In addition, it was reported that the long primary and secondary dendrites
that make up the main skeleton of the class IV arbors develop rather normally after γ-
tubulin depletion (Ori-McKenney et al., 2012). It remains unclear, however, whether γ-
TuRCs were efficiently depleted from the neurons in these experiments.

138

139

140

141

142

143

144

In this study, we have taken advantage of class IV arbor regrowth during pupal
development where we find that the effects of RNAi are stronger than in larvae. By
18h after pupal formation (APF) Class IV da neuron dendritic arbors are completely
removed in a process called pruning, leaving the neuronal cell body and axon intact
(Kuo et al., 2005; Williams and Truman, 2005). The class IV da neuron dendritic arbors
then have to regrow during pupal development to re-fill their receptive fields in
preparation for adulthood (Kuo et al., 2005; Satoh et al., 2012; Shimono et al., 2009).
Although microtubule regulation in da neurons is normally analysed in larvae, the
regrowth phase in pupae provides an opportunity to monitor the entirety of dendritic
arbor growth when RNAi constructs have been expressed for several days and when
there is less chance of a persistent maternal protein negating phenotypes.

145

146

147

148

149

150

151

152

153

154

155

156

Indeed, using the pupal system we show that γ-TuRCs are essential for many aspects
of class IV arbor development, including the growth and branching of primary and
secondary dendrites and for higher-order branching. These phenotypes are much less
severe in larvae, but can be detected when using the most efficient RNAi line against
γ-TuRCs. We also show that depletion of the Augmin complex predominantly affects
higher-order branching in both pupal and larval class IV neurons. In larvae, we observe
defects in microtubule polarity in distal secondary and tertiary dendrites and a
reduction in the frequency of microtubule growth within terminal dendrites after
Augmin depletion. This correlates with a reduction in the stability of these terminal
dendrites, which is thought to be microtubule dependent, but not in their initial growth,
which is actin dependent. Consistent with a specialised role for Augmin in higher-order
branching, it is expressed more strongly in larval class IV neurons than in class I
neurons, which have very few higher-order dendrites, and its depletion from class I

157

158

159

160

161

162

163

164

165

166

167

168
137

suggesting that the loss of terminal branches after γ-tubulin depletion may represent a failure in an as yet unidentified, Golgi outpost-independent, mode of microtubule nucleation. In addition, it was reported that the long primary and secondary dendrites that make up the main skeleton of the class IV arbors develop rather normally after γ-tubulin depletion (Ori-McKenney et al., 2012). It remains unclear, however, whether γ-TuRCs were efficiently depleted from the neurons in these experiments.

138

139

140

141

142

143

144

In this study, we have taken advantage of class IV arbor regrowth during pupal development where we find that the effects of RNAi are stronger than in larvae. By 18h after pupal formation (APF) Class IV da neuron dendritic arbors are completely removed in a process called pruning, leaving the neuronal cell body and axon intact (Kuo et al., 2005; Williams and Truman, 2005). The class IV da neuron dendritic arbors then have to regrow during pupal development to re-fill their receptive fields in preparation for adulthood (Kuo et al., 2005; Satoh et al., 2012; Shimono et al., 2009).

145

146

147

148

149

150

151

Although microtubule regulation in da neurons is normally analysed in larvae, the regrowth phase in pupae provides an opportunity to monitor the entirety of dendritic arbor growth when RNAi constructs have been expressed for several days and when there is less chance of a persistent maternal protein negating phenotypes.

152

153

154

155

156

Indeed, using the pupal system we show that γ-TuRCs are essential for many aspects of class IV arbor development, including the growth and branching of primary and secondary dendrites and for higher-order branching. These phenotypes are much less severe in larvae, but can be detected when using the most efficient RNAi line against γ-TuRCs. We also show that depletion of the Augmin complex predominantly affects higher-order branching in both pupal and larval class IV neurons. In larvae, we observe defects in microtubule polarity in distal secondary and tertiary dendrites and a reduction in the frequency of microtubule growth within terminal dendrites after Augmin depletion. This correlates with a reduction in the stability of these terminal dendrites, which is thought to be microtubule dependent, but not in their initial growth, which is actin dependent. Consistent with a specialised role for Augmin in higher-order branching, it is expressed more strongly in larval class IV neurons than in class I neurons, which have very few higher-order dendrites, and its depletion from class I

157

158

159

160

161

162

163

164

165

166

167

168
neurons has little effect on arbor morphology. Given the known role and mode of
operation of Augmin in other cell types, our findings suggest that Augmin mediates
sparsely distributed microtubule nucleation events throughout the dendritic arbor of
class IV da neurons that provide sufficient numbers of microtubules to stabilise the
nascent growth of higher-order dendrites. Our data also suggest that class IV and
class I da neurons use different modes of microtubule nucleation to support the
differential growth of their dendritic arbors.
169 neurons has little effect on arbor morphology. Given the known role and mode of
170 operation of Augmin in other cell types, our findings suggest that Augmin mediates
171 sparsely distributed microtubule nucleation events throughout the dendritic arbor of
172 class IV da neurons that provide sufficient numbers of microtubules to stabilise the
173 nascent growth of higher-order dendrites. Our data also suggest that class IV and
174 class I da neurons use different modes of microtubule nucleation to support the
175 differential growth of their dendritic arbors.
176 **Results**

177 **Depletion of γ-TuRCs strongly perturbs the development of pupal class IV da neurons**
178 We began by establishing imaging conditions for class IV da neurons during pupal
179 development. We chose to analyse v'ada class IV neurons as their dendritic arbor
180 development has been previously described (Yoong et al., 2020) and they remain free
181 from overlap by other fluorescent cells. Dendrites initially form at ~28-30h after pupae
182 formation (APF) and then undergo outgrowth and branching up until ~80h APF (Figure
183 1A). Similar to larval ddaC neurons, primary and secondary dendrites make up the
184 backbone of the arbor in pupae and higher-order branches help fill the dendritic field
185 (Figure S1). We tested the effect of depleting γ-TuRCs on the growth and development
186 of these arbors by expressing various UAS-controlled RNAi constructs in the presence
187 of UAS-Dicer2 using the class IV-specific driver ppk-Gal4. We chose to perform arbor
188 morphology analysis at ~72h APF because at this stage the arbors had grown to a
189 convenient size for analysis. Of all the γ-TuRC components, only γ-tubulin, Grip84
190 and Grip91, members of the γ-tubulin small complex (γ-TuSC), are essential for
191 viability and conserved throughout eukaryotes (Tovey and Conduit, 2018). We
192 therefore tested 2 independent RNAi lines for each of these essential γ-TuRC
193 components (Table S1). There are two γ-tubulin genes in *Drosophila*, a maternally
194 expressed form called γ-tubulin37C and a zygotically expressed form called γ-
195 tubulin23C. RNAi targeting γ-tubulin37C can therefore be used as a negative control
196 within da neurons (Feng et al., 2019; Mukherjee et al., 2020) and was used throughout
197 this study. For clarity, we have added the names of the Grip84 and Grip91 mammalian
198 homologues in superscript i.e. Grip84<sup>GCP2</sup> and Grip91<sup>GCP3</sup>.

199

200 For each condition, we measured both the number of dendrite tips and the arbor area
201 (defined by drawing a line around the perimeter of the dendritic arbor – hereafter
202 perimeter area – see Figure S1A and Methods section) at 72h APF. All but one RNAi
203 construct resulted in a significant reduction in tip number compared to the control, and
204 all but two resulted in a significant reduction in perimeter area (Figure 1B-D). As
205 expected, phenotypic strength varied, presumably due to differences in RNAi
206 efficiency, with the strongest reduction in both perimeter area and tip number resulting
207 from expressing grip91<sup>GCP3</sup>-RNAi1 (corresponding to VDRC KK library construct
176 **Results**

177 **Depletion of γ-TuRCs strongly perturbs the development of pupal class IV da neurons**
178 We began by establishing imaging conditions for class IV da neurons during pupal
179 development. We chose to analyse v'ada class IV neurons as their dendritic arbor
180 development has been previously described (Yoong et al., 2020) and they remain free
181 from overlap by other fluorescent cells. Dendrites initially form at ~28-30h after pupae
182 formation (APF) and then undergo outgrowth and branching up until ~80h APF (Figure
183 1A). Similar to larval ddaC neurons, primary and secondary dendrites make up the
184 backbone of the arbor in pupae and higher-order branches help fill the dendritic field
185 (Figure S1). We tested the effect of depleting γ-TuRCs on the growth and development
186 of these arbors by expressing various UAS-controlled RNAi constructs in the presence
187 of UAS-Dicer2 using the class IV-specific driver ppk-Gal4. We chose to perform arbor
188 morphology analysis at ~72h APF because at this stage the arbors had grown to a
189 convenient size for analysis. Of all the γ-TuRC components, only γ-tubulin, Grip84
190 and Grip91, members of the γ-tubulin small complex (γ-TuSC), are essential for
191 viability and conserved throughout eukaryotes (Tovey and Conduit, 2018). We
192 therefore tested 2 independent RNAi lines for each of these essential γ-TuRC
193 components (Table S1). There are two γ-tubulin genes in *Drosophila*, a maternally
194 expressed form called γ-tubulin37C and a zygotically expressed form called γ-
195 tubulin23C. RNAi targeting γ-tubulin37C can therefore be used as a negative control
196 within da neurons (Feng et al., 2019; Mukherjee et al., 2020) and was used throughout
197 this study. For clarity, we have added the names of the Grip84 and Grip91 mammalian
198 homologues in superscript i.e. Grip84<sup>GCP2</sup> and Grip91<sup>GCP3</sup>.

199

200 For each condition, we measured both the number of dendrite tips and the arbor area
201 (defined by drawing a line around the perimeter of the dendritic arbor – hereafter
202 perimeter area – see Figure S1A and Methods section) at 72h APF. All but one RNAi
203 construct resulted in a significant reduction in tip number compared to the control, and
204 all but two resulted in a significant reduction in perimeter area (Figure 1B-D). As
205 expected, phenotypic strength varied, presumably due to differences in RNAi
206 efficiency, with the strongest reduction in both perimeter area and tip number resulting
207 from expressing grip91<sup>GCP3</sup>-RNAi1 (corresponding to VDRC KK library construct
104667) and the second strongest resulting from expressing γ-tubulin23C-RNAi1, which was previously used when assessing phenotypes in larvae (Ori-McKenney et al., 2012) (Figure 1B-D). Although we cannot rule out that the stronger phenotype observed with grip91<sup>GCP3</sup>-RNAi1 is due to unintended depletion of another gene, grip91<sup>GCP3</sup>-RNAi1 has no predicted off-targets (VDRC website). Moreover, we confirmed by RT-qPCR on wing disc samples that grip91<sup>GCP3</sup>-RNAi1 was able to deplete grip91 mRNA (Figure S2) and that expression in class I da neurons strongly reduced the presence of endogenously-tagged γ-tubulin-GFP at Golgi mini-stacks within the soma (Figure S3). Thus, expressing grip91<sup>GCP3</sup>-RNAi1 has a strong effect on γ-TuRCs in da neurons.

218

We therefore further analysed pupal class IV neurons expressing grip91<sup>GCP3</sup>-RNAi. Strikingly, compared to controls, there was a ~75% and ~77% reduction in dendritic tips and perimeter area at 72h APF, respectively, when expressing grip91<sup>GCP3</sup>-RNAi (Figure 1B-D). The length of the longest primary dendrite that extends from the soma was reduced by ~55% (Figure 1E) and the combined length of primary and secondary dendrites was reduced by ~73% (Figure 1F). There was also a strong reduction in the combined total number of primary and secondary branchpoints (Figure 1G). Nevertheless, these major branchpoints occurred more frequently per dendrite length in grip91<sup>GCP3</sup>-RNAi neurons, suggesting that while the primary and secondary dendrites of grip91<sup>GCP3</sup>-RNAi neurons struggle to both grow (Figure 1F) and branch (Figure 1G) they perhaps struggle more to grow than to branch. Taking the data together, we conclude that γ-TuRCs are required for primary and secondary dendrite growth and branching and for higher-order branching.

219

220

221

222

223

224

225

226

227

228

229

230

231

232

233

234

### Depletion of γ-TuRCs perturbs the development of larval class IV da neurons, albeit more mildly than in pupal neurons

235

236

237

238

239

We considered that the phenotypes we observed in pupal neurons were stronger than those previously observed in larval neurons because of a maternal contribution of γ-TuRCs (provided by the mother to the embryo) that persists during larval stages. This possibility had already been suggested by others (Ori-McKenney et al., 2012) and is supported by our unpublished observations that γ-tubulin23c null mutant flies can
104667) and the second strongest resulting from expressing γ-tubulin23C-RNAi1, which was previously used when assessing phenotypes in larvae (Ori-McKenney et al., 2012) (Figure 1B-D). Although we cannot rule out that the stronger phenotype observed with grip91<sup>GCP3</sup>-RNAi1 is due to unintended depletion of another gene, grip91<sup>GCP3</sup>-RNAi1 has no predicted off-targets (VDRC website). Moreover, we confirmed by RT-qPCR on wing disc samples that grip91<sup>GCP3</sup>-RNAi1 was able to deplete grip91 mRNA (Figure S2) and that expression in class I da neurons strongly reduced the presence of endogenously-tagged γ-tubulin-GFP at Golgi mini-stacks within the soma (Figure S3). Thus, expressing grip91<sup>GCP3</sup>-RNAi1 has a strong effect on γ-TuRCs in da neurons.

218

We therefore further analysed pupal class IV neurons expressing grip91<sup>GCP3</sup>-RNAi. Strikingly, compared to controls, there was a ~75% and ~77% reduction in dendritic tips and perimeter area at 72h APF, respectively, when expressing grip91<sup>GCP3</sup>-RNAi (Figure 1B-D). The length of the longest primary dendrite that extends from the soma was reduced by ~55% (Figure 1E) and the combined length of primary and secondary dendrites was reduced by ~73% (Figure 1F). There was also a strong reduction in the combined total number of primary and secondary branchpoints (Figure 1G). Nevertheless, these major branchpoints occurred more frequently per dendrite length in grip91<sup>GCP3</sup>-RNAi neurons, suggesting that while the primary and secondary dendrites of grip91<sup>GCP3</sup>-RNAi neurons struggle to both grow (Figure 1F) and branch (Figure 1G) they perhaps struggle more to grow than to branch. Taking the data together, we conclude that γ-TuRCs are required for primary and secondary dendrite growth and branching and for higher-order branching.

219

220

221

222

223

224

225

226

227

228

229

230

231

232

233

234

### Depletion of γ-TuRCs perturbs the development of larval class IV da neurons, albeit more mildly than in pupal neurons

235

236

237

238

239

We considered that the phenotypes we observed in pupal neurons were stronger than those previously observed in larval neurons because of a maternal contribution of γ-TuRCs (provided by the mother to the embryo) that persists during larval stages. This possibility had already been suggested by others (Ori-McKenney et al., 2012) and is supported by our unpublished observations that γ-tubulin23c null mutant flies can
240

survive until late larval/early pupal stages. RNAi expression may also be relatively low
during the early stages of arbor development in larvae, and the previously observed
phenotypes in larvae may have been stronger if the most efficient γ-TuRC RNAi line
had been used.

241

242

243

244

245

To examine the effect of depleting γ-TuRCs on larval class IV neurons, we expressed grip91<sup>GCP3</sup>-RNAi with UAS-Dicer2 and examined class IV ddaC arbors at 72h and 96h after egg laying (AEL). The growth of class IV da neurons has been well documented (Gao et al., 1999; Parrish et al., 2009; Shree et al., 2022). Dendrite growth begins in late embryos at around 16h AEL. During the first and second larval instar stages, from 24h to 72h AEL, the dendritic arbors grow faster than the hemisegment boundaries until they fill the hemisegments at ~72h AEL. The arbors then grow at the same rate as the hemisegments, termed scaling growth, until 96h AEL. During 96h-120h AEL, the wandering third instar stage, the arbors occupy the same area but continue to add internal branches, maximising their density and coverage of the dendritic field. We found that there was no significant difference in perimeter area between control and grip91<sup>GCP3</sup>-RNAi arbors at 72h AEL (Figure 2A,B), suggesting that the growth of the main primary and secondary dendrites had been unaffected. There was, however, a ~40% reduction in dendrite tips (Figure 2A,B), showing that higher-order branches failed to properly form or be maintained. At 96h AEL, while the control arbors had scaled with segment growth and increased their perimeter area by ~15%, grip91<sup>GCP3</sup>-RNAi arbors failed to scale and remained at a similar perimeter area to the arbors at 72h AEL, while also having a ~35% reduction in dendrite tips (Figure 2C,D). The failure to scale was made obvious by the appearance of large gaps between adjacent grip91<sup>GCP3</sup>-RNAi neurons (orange arrowheads, Figure 2C). Thus, expressing grip91<sup>GCP3</sup>-RNAi perturbs the growth and/or stability of higher-order dendrites at both 72h and 96h AEL, but takes effect on the outward growth of primary and secondary dendrites only after 72h AEL, during scaling growth. These data are consistent with an inefficiency of RNAi during early dendritic arbor growth, particularly concerning primary and secondary dendrite growth.

246

247

248

249

250

251

252

253

254

255

256

257

258

259

260

261

262

263

264

265

266

267

268

269

270
240

survive until late larval/early pupal stages. RNAi expression may also be relatively low
during the early stages of arbor development in larvae, and the previously observed
phenotypes in larvae may have been stronger if the most efficient γ-TuRC RNAi line
had been used.

241

242

243

244

245

To examine the effect of depleting γ-TuRCs on larval class IV neurons, we expressed grip91<sup>GCP3</sup>-RNAi with UAS-Dicer2 and examined class IV ddaC arbors at 72h and 96h after egg laying (AEL). The growth of class IV da neurons has been well documented (Gao et al., 1999; Parrish et al., 2009; Shree et al., 2022). Dendrite growth begins in late embryos at around 16h AEL. During the first and second larval instar stages, from 24h to 72h AEL, the dendritic arbors grow faster than the hemisegment boundaries until they fill the hemisegments at ~72h AEL. The arbors then grow at the same rate as the hemisegments, termed scaling growth, until 96h AEL. During 96h-120h AEL, the wandering third instar stage, the arbors occupy the same area but continue to add internal branches, maximising their density and coverage of the dendritic field. We found that there was no significant difference in perimeter area between control and grip91<sup>GCP3</sup>-RNAi arbors at 72h AEL (Figure 2A,B), suggesting that the growth of the main primary and secondary dendrites had been unaffected. There was, however, a ~40% reduction in dendrite tips (Figure 2A,B), showing that higher-order branches failed to properly form or be maintained. At 96h AEL, while the control arbors had scaled with segment growth and increased their perimeter area by ~15%, grip91<sup>GCP3</sup>-RNAi arbors failed to scale and remained at a similar perimeter area to the arbors at 72h AEL, while also having a ~35% reduction in dendrite tips (Figure 2C,D). The failure to scale was made obvious by the appearance of large gaps between adjacent grip91<sup>GCP3</sup>-RNAi neurons (orange arrowheads, Figure 2C). Thus, expressing grip91<sup>GCP3</sup>-RNAi perturbs the growth and/or stability of higher-order dendrites at both 72h and 96h AEL, but takes effect on the outward growth of primary and secondary dendrites only after 72h AEL, during scaling growth. These data are consistent with an inefficiency of RNAi during early dendritic arbor growth, particularly concerning primary and secondary dendrite growth.

246

247

248

249

250

251

252

253

254

255

256

257

258

259

260

261

262

263

264

265

266

267

268

269

270
271

Collectively, our data from pupal and larval neurons show that γ-TuRCs are required
for the proper growth and development of class IV dendritic arbors, with their depletion
affecting primary and secondary dendrite growth (during later larval stages and
throughout pupal development) and the growth/stability of the more dynamic higher-
order branches.

272

273

274

275

276

277

Depletion of Augmin perturbs dendritic branching of pupal class IV da neurons

278

279

280

281

282

283

284

285

286

287

288

289

290

291

292

293

294

295

296

297

298

299

300

301

302

Collating the finding above that γ-TuRC depletion perturbs higher-order branching in class IV neurons with the previous observation that distal Golgi outposts lack γ-TuRCs (Mukherjee et al., 2020), we considered which other modes of γ-TuRC-mediated microtubule nucleation may be involved in regulating higher-order branching. The Augmin complex was a good candidate, as it recruits γ-TuRCs to the sides of pre-existing microtubules from which new daughter microtubules are nucleated. This mode of nucleation could in theory occur throughout the arbor. Moreover, these nucleation events can be angled up to ~60° depending on the organism and cell type (~36° in *Drosophila* S2 cells (Verma and Maresca, 2019)), which could provide a mechanism to promote microtubule growth into nascent higher-order dendrites.

We first tested whether Augmin components were expressed in class IV da neurons.
We crossed a UAS-myr-GFP line (GFP targeted to membrane) to a publicly available
enhancer trap line where Gal4 had been inserted into the gene region of the Augmin
component Dgt5 (BL77584). We observed GFP signal within 35/35 larval class IV
ddaC neurons from 6 different animals (Figure S4), consistent with the expression of
the Augmin complex. We then examined the effects on pupal neurons of expressing
RNAi constructs targeting Augmin. Augmin comprises 8 different proteins and its
ability to recruit γ-TuRCs to spindle microtubules depends on the γ-TuRC adapter
protein Grip71 (homologue of NEDD1). We therefore expressed different RNAi lines
against various Augmin components (dgt2, dgt3, dgt4, dgt5) or grip71<sup>NEDD1</sup> (Table S2).
We confirmed by PCR that none of the KK lines contained the aberrant 40D insertion
(data not shown). All but one of the eight RNAi lines, including both grip71<sup>NEDD1</sup>-RNAi
lines, caused a significant reduction in dendrite tip number and perimeter area (Figure
3A-C), consistent with a role for Augmin in mediating class IV dendritic arbor growth.
271

Collectively, our data from pupal and larval neurons show that γ-TuRCs are required for the proper growth and development of class IV dendritic arbors, with their depletion affecting primary and secondary dendrite growth (during later larval stages and throughout pupal development) and the growth/stability of the more dynamic higher-order branches.

272

273

274

275

276

277

### Depletion of Augmin perturbs dendritic branching of pupal class IV da neurons

278

279

280

281

282

283

284

285

286

287

Collating the finding above that γ-TuRC depletion perturbs higher-order branching in class IV neurons with the previous observation that distal Golgi outposts lack γ-TuRCs (Mukherjee et al., 2020), we considered which other modes of γ-TuRC-mediated microtubule nucleation may be involved in regulating higher-order branching. The Augmin complex was a good candidate, as it recruits γ-TuRCs to the sides of pre-existing microtubules from which new daughter microtubules are nucleated. This mode of nucleation could in theory occur throughout the arbor. Moreover, these nucleation events can be angled up to ~60° depending on the organism and cell type (~36° in *Drosophila* S2 cells (Verma and Maresca, 2019)), which could provide a mechanism to promote microtubule growth into nascent higher-order dendrites.

288

289

290

291

292

293

294

295

296

297

298

We first tested whether Augmin components were expressed in class IV da neurons. We crossed a UAS-myr-GFP line (GFP targeted to membrane) to a publicly available enhancer trap line where Gal4 had been inserted into the gene region of the Augmin component Dgt5 (BL77584). We observed GFP signal within 35/35 larval class IV ddaC neurons from 6 different animals (Figure S4), consistent with the expression of the Augmin complex. We then examined the effects on pupal neurons of expressing RNAi constructs targeting Augmin. Augmin comprises 8 different proteins and its ability to recruit γ-TuRCs to spindle microtubules depends on the γ-TuRC adapter protein Grip71 (homologue of NEDD1). We therefore expressed different RNAi lines against various Augmin components (dgt2, dgt3, dgt4, dgt5) or grip71<sup>NEDD1</sup> (Table S2).

299

300

301

302

We confirmed by PCR that none of the KK lines contained the aberrant 40D insertion (data not shown). All but one of the eight RNAi lines, including both grip71<sup>NEDD1</sup>-RNAi lines, caused a significant reduction in dendrite tip number and perimeter area (Figure 3A-C), consistent with a role for Augmin in mediating class IV dendritic arbor growth.
The strongest effect came when expressing Dgt4-RNAi, which resulted in a ~52% reduction in perimeter area (Figure 3B) and a ~50% reduction in tip number (Figure 3C), and was capable of depleting dgt4 mRNA when expressed in wing discs (Figure S2). In contrast to the effects after γ-TuRC depletion, however, there was no significant reduction in the length of the longest primary dendrite (Figure 3D), and there was only a ~27% reduction in the total length of primary and secondary branches (compared to a ~73% reduction in grip91<sup>GCP3</sup>-RNAi neurons) (Figure 3E). Moreover, there was no effect on the number of primary and secondary branchpoints (Figure 3F), meaning that the length of primary and secondary dendrite per branchpoint was reduced slightly (Figure 3G). Thus, while there is some effect on primary and secondary dendrite growth, much of the reduction in peripheral arbour area and tip number after Augmin depletion appears to be due to defects in higher-order branching. Depleting the γ-TuRC component Grip71<sup>NEDD1</sup> produced very similar phenotypes (Figure 3B-G), suggesting that Grip71<sup>NEDD1</sup> functions specifically in the Augmin pathway in these neurons, rather than acting as a general γ-TuRC targeting.

318

319 Depletion of Augmin perturbs dendritic branching within larval class IV da neurons

We next tested the effect of Augmin depletion within larval class IV neurons. While RNAi may be less efficient than in pupae, the dynamics of microtubules are well characterised in larval neurons (Ori-McKenney et al., 2012; Sears and Broihier, 2016), allowing us to carefully examine the effect of Augmin depletion on microtubule polarity and growth. Moreover, we could compare effects between class I and class IV neurons, which have very different dendritic arbor morphologies. We expressed Dgt4-RNAi and UAS-Dicer2 within class IV da neurons using the ppk-gal4 driver and assessed dendritic arbor morphology at 96h AEL. Compared to controls, there was a ~17% and a ~13% reduction in dendrite tip number and perimeter area, respectively (Figure 4A,B). Intriguingly, however, the peripheral area was similar to controls 24h later, at 120h AEL, just before pupation, despite there still being a ~22% reduction in dendrite tips (Figure 4C,D). We also measured a ~32% reduction in average terminal branch length in Dgt4-RNAi neurons at 96h AEL (Figure 4E), with a downward shift in the distribution of terminal dendrite length (Figure 4F). These data are consistent with Augmin having a more important role in higher-order branching than in the growth of
The strongest effect came when expressing Dgt4-RNAi, which resulted in a ~52% reduction in perimeter area (Figure 3B) and a ~50% reduction in tip number (Figure 3C), and was capable of depleting dgt4 mRNA when expressed in wing discs (Figure S2). In contrast to the effects after γ-TuRC depletion, however, there was no significant reduction in the length of the longest primary dendrite (Figure 3D), and there was only a ~27% reduction in the total length of primary and secondary branches (compared to a ~73% reduction in grip91<sup>GCP3</sup>-RNAi neurons) (Figure 3E). Moreover, there was no effect on the number of primary and secondary branchpoints (Figure 3F), meaning that the length of primary and secondary dendrite per branchpoint was reduced slightly (Figure 3G). Thus, while there is some effect on primary and secondary dendrite growth, much of the reduction in peripheral arbour area and tip number after Augmin depletion appears to be due to defects in higher-order branching. Depleting the γ-TuRC component Grip71<sup>NEDD1</sup> produced very similar phenotypes (Figure 3B-G), suggesting that Grip71<sup>NEDD1</sup> functions specifically in the Augmin pathway in these neurons, rather than acting as a general γ-TuRC targeting.

318

319 Depletion of Augmin perturbs dendritic branching within larval class IV da neurons

320 We next tested the effect of Augmin depletion within larval class IV neurons. While 321 RNAi may be less efficient than in pupae, the dynamics of microtubules are well 322 characterised in larval neurons (Ori-McKenney et al., 2012; Sears and Broihier, 2016), 323 allowing us to carefully examine the effect of Augmin depletion on microtubule polarity 324 and growth. Moreover, we could compare effects between class I and class IV 325 neurons, which have very different dendritic arbor morphologies. We expressed Dgt4- 326 RNAi and UAS-Dicer2 within class IV da neurons using the ppk-gal4 driver and 327 assessed dendritic arbor morphology at 96h AEL. Compared to controls, there was a 328 ~17% and a ~13% reduction in dendrite tip number and perimeter area, respectively 329 (Figure 4A,B). Intriguingly, however, the peripheral area was similar to controls 24h 330 later, at 120h AEL, just before pupation, despite there still being a ~22% reduction in 331 dendrite tips (Figure 4C,D). We also measured a ~32% reduction in average terminal 332 branch length in Dgt4-RNAi neurons at 96h AEL (Figure 4E), with a downward shift in 333 the distribution of terminal dendrite length (Figure 4F). These data are consistent with 334 Augmin having a more important role in higher-order branching than in the growth of
335

primary and secondary dendrites, and this was made apparent by the presence of
large areas void of higher-order dendrites within Dgt4-RNAi neurons (compare insets
in Figure 4A and 4C).

336

337

338

339

### Depletion of Augmin perturbs microtubule polarity and frequency within the distal regions larval class IV da neurons

340

341

To determine whether the reduction in higher-order branching of class IV da neurons correlates with defects in microtubule dynamics, we expressed the plus-end tracking protein EB1-GFP in either control-RNAi or Dgt4-RNAi neurons (see Movie S1 and Movie S2). EB1-GFP marks the growing ends of microtubules and is commonly used to determine microtubule polarity and new microtubule growth events. In class IV da neurons, the direction of EB1-GFP comet growth correlates with dendrite length, with comets becoming progressively more anterograde (growing away from the soma) with reduced branch length, which roughly correlates with dendrite order (primary, secondary, tertiary, terminal) (Ori-McKenney et al., 2012). Indeed, when imaging the long primary dendrites close to the soma we found that 213/216 (98.6%) EB1-GFP comets were retrograde in control-RNAi neurons. Consistent with Augmin not playing a role in the growth of primary dendrites, there was no significant decrease in the proportion of retrograde comets in these primary dendrites (192/201 (95.5%); p=0.079) (Figure 4G).

342

343

344

345

346

347

348

349

350

351

352

353

354

355

356

When filming the more distal regions of control arbors, however, we could identify differences in comet polarity and frequency. In secondary and tertiary dendrites (see Figure S1B for how dendrite order was established), there was a significant reduction in the proportion of retrograde comets in Dgt4-RNAi neurons: 73.8% (220/298) and 60.9% (380/622) of comets were retrograde in control-RNAi or Dgt4-RNAi neurons, respectively (p<0.001) (Figure 4G). Thus, Augmin depletion can affect microtubule polarity in *Drosophila* dendrites, as it does in mammalian axons (Cunha-Ferreira et al., 2018; Sánchez-Huertas et al., 2016). In contrast to secondary and tertiary dendrites, however, a higher frequency of retrograde comets was observed in the terminal dendrites of Dgt4-RNAi neurons compared to control neurons, although this was not statistically significant (Figure 4G). When quantifying EB1-GFP comet frequency, we

357

358

359

360

361

362

363

364

365

366
335

primary and secondary dendrites, and this was made apparent by the presence of
large areas void of higher-order dendrites within Dgt4-RNAi neurons (compare insets
in Figure 4A and 4C).

336

337

338

339

### Depletion of Augmin perturbs microtubule polarity and frequency within the distal regions larval class IV da neurons

340

341

To determine whether the reduction in higher-order branching of class IV da neurons correlates with defects in microtubule dynamics, we expressed the plus-end tracking protein EB1-GFP in either control-RNAi or Dgt4-RNAi neurons (see Movie S1 and Movie S2). EB1-GFP marks the growing ends of microtubules and is commonly used to determine microtubule polarity and new microtubule growth events. In class IV da neurons, the direction of EB1-GFP comet growth correlates with dendrite length, with comets becoming progressively more anterograde (growing away from the soma) with reduced branch length, which roughly correlates with dendrite order (primary, secondary, tertiary, terminal) (Ori-McKenney et al., 2012). Indeed, when imaging the long primary dendrites close to the soma we found that 213/216 (98.6%) EB1-GFP comets were retrograde in control-RNAi neurons. Consistent with Augmin not playing a role in the growth of primary dendrites, there was no significant decrease in the proportion of retrograde comets in these primary dendrites (192/201 (95.5%); p=0.079) (Figure 4G).

342

343

344

345

346

347

348

349

350

351

352

353

354

355

356

When filming the more distal regions of control arbors, however, we could identify differences in comet polarity and frequency. In secondary and tertiary dendrites (see Figure S1B for how dendrite order was established), there was a significant reduction in the proportion of retrograde comets in Dgt4-RNAi neurons: 73.8% (220/298) and 60.9% (380/622) of comets were retrograde in control-RNAi or Dgt4-RNAi neurons, respectively (p<0.001) (Figure 4G). Thus, Augmin depletion can affect microtubule polarity in *Drosophila* dendrites, as it does in mammalian axons (Cunha-Ferreira et al., 2018; Sánchez-Huertas et al., 2016). In contrast to secondary and tertiary dendrites, however, a higher frequency of retrograde comets was observed in the terminal dendrites of Dgt4-RNAi neurons compared to control neurons, although this was not statistically significant (Figure 4G). When quantifying EB1-GFP comet frequency, we

357

358

359

360

361

362

363

364

365

366
found, somewhat surprisingly, that distal secondary and tertiary dendrites displayed a ~24% increase in EB1-GFP comets (Figure 4H). In stark contrast, however, there was a ~44% reduction in EB1-GFP comet frequency in distal terminal dendrites (Figure 4H). The increase in comet frequency in distal secondary and tertiary dendrites could be due to the induction of neuronal stress, as is the case after the depletion of other microtubule regulators (Feng et al., 2019). Alternatively, the reduction in microtubule growth in terminal dendrites may increase the availability of tubulin dimers for microtubule growth in distal secondary and tertiary dendrites. In either case, we conclude that Augmin is required to maintain proper microtubule polarity in distal secondary and tertiary dendrites and to promote anterograde microtubule growth within terminal dendrites.

### Terminal dendrites are destabilised in Dgt4-RNAi neurons

We next examined the dynamics of terminal dendrites in control-RNAi and Dgt4-RNAi neurons. It is known that terminal dendrites are highly dynamic (Shree et al., 2022), forming in an actin-dependent manner (Nithianandam and Chien, 2018; Stürner et al., 2019) and then either disappearing, remaining stable or extending their growth. Moreover, stability correlates with the presence of anterograde EB1-GFP comets (Ori-McKenney et al., 2012; Sears and Broihier, 2016). We filmed distal regions of control-RNAi and Dgt4-RNAi neurons expressing a fluorescent membrane marker for 30 mins and found no significant difference in the rate at which new terminal dendrites formed (Figure 4I), consistent with Augmin not being involved in actin remodelling. Strikingly, however, we found that the percentage of initial terminal dendrites that disappeared by the end of the movie increased from 23% in control-RNAi neurons to 40% in Dgt4-RNAi neurons (Figure 4J). Thus, Augmin's ability to promote anterograde microtubule growth within terminal dendrites appears to help stabilise these terminal dendrites, which presumably promotes higher-order branching.

### Augmin is largely dispensable for the development of larval class I da neurons

Given that larval class I da neurons have only few higher-order dendrites and mainly comprise primary and secondary dendrites, we hypothesised that Augmin may be largely dispensable for class I arbor development. Consistent with this, the dgt5 Gal4
found, somewhat surprisingly, that distal secondary and tertiary dendrites displayed a ~24% increase in EB1-GFP comets (Figure 4H). In stark contrast, however, there was a ~44% reduction in EB1-GFP comet frequency in distal terminal dendrites (Figure 4H). The increase in comet frequency in distal secondary and tertiary dendrites could be due to the induction of neuronal stress, as is the case after the depletion of other microtubule regulators (Feng et al., 2019). Alternatively, the reduction in microtubule growth in terminal dendrites may increase the availability of tubulin dimers for microtubule growth in distal secondary and tertiary dendrites. In either case, we conclude that Augmin is required to maintain proper microtubule polarity in distal secondary and tertiary dendrites and to promote anterograde microtubule growth within terminal dendrites.

### Terminal dendrites are destabilised in Dgt4-RNAi neurons

We next examined the dynamics of terminal dendrites in control-RNAi and Dgt4-RNAi neurons. It is known that terminal dendrites are highly dynamic (Shree et al., 2022), forming in an actin-dependent manner (Nithianandam and Chien, 2018; Stürner et al., 2019) and then either disappearing, remaining stable or extending their growth. Moreover, stability correlates with the presence of anterograde EB1-GFP comets (Ori-McKenney et al., 2012; Sears and Broihier, 2016). We filmed distal regions of control-RNAi and Dgt4-RNAi neurons expressing a fluorescent membrane marker for 30 mins and found no significant difference in the rate at which new terminal dendrites formed (Figure 4I), consistent with Augmin not being involved in actin remodelling. Strikingly, however, we found that the percentage of initial terminal dendrites that disappeared by the end of the movie increased from 23% in control-RNAi neurons to 40% in Dgt4-RNAi neurons (Figure 4J). Thus, Augmin's ability to promote anterograde microtubule growth within terminal dendrites appears to help stabilise these terminal dendrites, which presumably promotes higher-order branching.

### Augmin is largely dispensable for the development of larval class I da neurons

Given that larval class I da neurons have only few higher-order dendrites and mainly comprise primary and secondary dendrites, we hypothesised that Augmin may be largely dispensable for class I arbor development. Consistent with this, the dgt5 Gal4
enhancer trap was expressed less strongly in class I ddaE neurons than in class IV ddaC neurons (Figure S4C). Moreover, expressing Dgt4-RNAi in class I neurons had little effect on dendritic arbor morphology. In Dgt4-RNAi neurons, there was only a ~8.4% and ~11.4% reduction in total arbor length at 96h AEL (Figure 5A,B) and 120h AEL (Figure 5C,D), respectively, showing that Augmin is largely dispensable for the proper growth of class I da neuron primary and secondary dendrites (similar to class IV da neurons). There were also no significant changes in dendrite tip number at either 96h (Figure 5A,B) or 120h (Figure 5C,D) AEL. This is consistent with a previous observation that embryonic class I ddaE neurons within augmin mutant flies also show no difference in dendrite tip number compared to controls (Yalgin et al., 2015). In control-RNAi and Dgt4-RNAi larval class I neurons, we measured an average of ~36 dendrite tips at 120h AEL, while Yalgin et al. measured an average of ~20 dendrite tips during stage 17 of embryogenesis (just before hatching into larvae), ~96 hours earlier. Thus, class I ddaE neurons add, on average, ~16 new dendrite branches during ~96 hours of larval development and this does not require Augmin.
enhancer trap was expressed less strongly in class I ddaE neurons than in class IV ddaC neurons (Figure S4C). Moreover, expressing Dgt4-RNAi in class I neurons had little effect on dendritic arbor morphology. In Dgt4-RNAi neurons, there was only a ~8.4% and ~11.4% reduction in total arbor length at 96h AEL (Figure 5A,B) and 120h AEL (Figure 5C,D), respectively, showing that Augmin is largely dispensable for the proper growth of class I da neuron primary and secondary dendrites (similar to class IV da neurons). There were also no significant changes in dendrite tip number at either 96h (Figure 5A,B) or 120h (Figure 5C,D) AEL. This is consistent with a previous observation that embryonic class I ddaE neurons within augmin mutant flies also show no difference in dendrite tip number compared to controls (Yalgin et al., 2015). In control-RNAi and Dgt4-RNAi larval class I neurons, we measured an average of ~36 dendrite tips at 120h AEL, while Yalgin et al. measured an average of ~20 dendrite tips during stage 17 of embryogenesis (just before hatching into larvae), ~96 hours earlier. Thus, class I ddaE neurons add, on average, ~16 new dendrite branches during ~96 hours of larval development and this does not require Augmin.
414 **Discussion**

415 Our data show that γ-TuRCs and the Augmin complex are required for the growth and
416 development of highly-branched class IV dendritic arbors. γ-TuRCs are required for
417 all aspects of dendritic arbor development, while Augmin appears to be required
418 predominantly for higher-order branching. Consistent with this, Augmin depletion has
419 little effect in class I neurons, which have only few higher-order dendrites. Thus, while
420 γ-TuRCs are generally required for dendritic arbor development, the Augmin complex
421 may be a key determinant of dendritic arbor morphology, promoting the elaboration of
422 dendritic arbors.

423

424 While we cannot exclude the possibility that Augmin functions independently of γ-
425 TuRCs in da neurons, a γ-TuRC-independent function of Augmin has not been
426 reported to our knowledge. Given the effects we observe on microtubule polarity and
427 frequency after Augmin depletion, our data are fully consistent with Augmin functioning
428 to recruit γ-TuRCs to the sides of microtubules to mediate microtubule nucleation
429 events, as occurs in cycling cells (Song et al., 2018). We propose that these nucleation
430 events occur throughout the arbor and lead to the anterograde growth of microtubules
431 within terminal dendrites, which in turn help to stabilise the nascent growth of these
432 dendrites.

433

434 Our finding that Augmin is required for higher-order branching in class IV neurons
435 helps make sense of prior literature. Higher-order branching in larvae was shown to
436 be reduced after γ-tubulin depletion (Ori-McKenney et al., 2012). This was attributed
437 to microtubule nucleation from γ-TuRCs at Golgi outposts (Ori-McKenney et al., 2012),
438 but it was later shown that most Golgi outposts, including those positioned in the distal
439 regions of the arbor, do not recruit detectable levels of γ-tubulin-GFP (Mukherjee et
440 al., 2020). Based on our current data, we propose that γ-TuRCs are instead recruited
441 via Augmin to the sides of pre-existing microtubules. There are several non-mutually
442 exclusive possibilities for how microtubules nucleated from these γ-TuRCs may
443 ultimately grow within terminal dendrites: 1) Augmin-bound γ-TuRCs may nucleate
444 microtubules at the base of nascent branchpoints with the daughter microtubule
445 growing directly into the nascent dendrite (Figure 6A). Recruitment of Augmin to the
414 Discussion

415 Our data show that γ-TuRCs and the Augmin complex are required for the growth and
416 development of highly-branched class IV dendritic arbors. γ-TuRCs are required for
417 all aspects of dendritic arbor development, while Augmin appears to be required
418 predominantly for higher-order branching. Consistent with this, Augmin depletion has
419 little effect in class I neurons, which have only few higher-order dendrites. Thus, while
420 γ-TuRCs are generally required for dendritic arbor development, the Augmin complex
421 may be a key determinant of dendritic arbor morphology, promoting the elaboration of
422 dendritic arbors.
423

423

424 While we cannot exclude the possibility that Augmin functions independently of γ-
425 TuRCs in da neurons, a γ-TuRC-independent function of Augmin has not been
426 reported to our knowledge. Given the effects we observe on microtubule polarity and
427 frequency after Augmin depletion, our data are fully consistent with Augmin functioning
428 to recruit γ-TuRCs to the sides of microtubules to mediate microtubule nucleation
429 events, as occurs in cycling cells (Song et al., 2018). We propose that these nucleation
430 events occur throughout the arbor and lead to the anterograde growth of microtubules
431 within terminal dendrites, which in turn help to stabilise the nascent growth of these
432 dendrites.
433

433

434 Our finding that Augmin is required for higher-order branching in class IV neurons
435 helps make sense of prior literature. Higher-order branching in larvae was shown to
436 be reduced after γ-tubulin depletion (Ori-McKenney et al., 2012). This was attributed
437 to microtubule nucleation from γ-TuRCs at Golgi outposts (Ori-McKenney et al., 2012),
438 but it was later shown that most Golgi outposts, including those positioned in the distal
439 regions of the arbor, do not recruit detectable levels of γ-tubulin-GFP (Mukherjee et
440 al., 2020). Based on our current data, we propose that γ-TuRCs are instead recruited
441 via Augmin to the sides of pre-existing microtubules. There are several non-mutually
442 exclusive possibilities for how microtubules nucleated from these γ-TuRCs may
443 ultimately grow within terminal dendrites: 1) Augmin-bound γ-TuRCs may nucleate
444 microtubules at the base of nascent branchpoints with the daughter microtubule
445 growing directly into the nascent dendrite (Figure 6A). Recruitment of Augmin to the
446

branchpoint could be stochastic, helping to explain why many nascent dendrites retract and disappear, or it could be influenced by the presence of other regulators of branching, such as Arp2/3 (Stürner et al., 2019); 2) Augmin could amplify microtubules throughout the arbor, increasing the possibility that microtubules turn into nascent dendrites (Figure 6B), although our EB1-GFP data shows that microtubule growth is actually increased in secondary and tertiary dendrites after Augmin depletion; 3) Augmin may amplify anterograde microtubules already present within nascent dendrites (Figure 6C). An alternative possibility is that Augmin helps maintain correct microtubule polarity to ensure the proper delivery of material needed for new branching events (Figure 6D). Future studies will be required to elucidate the exact molecular mechanism underlying Augmin's role in dendrite branching.

447

448

449

450

451

452

453

454

455

456

457

458

459

460

461

462

463

464

465

466

467

468

469

470

471

472

473

474

475

476

477

Intriguingly, we find that Grip7¹<sup>NEDD1</sup> depletion results in similar phenotypes to Augmin depletion. However, both these phenotypes are weaker than those resulting from γ-TuRC depletion. Firstly, this suggests that the γ-TuRC protein Grip71<sup>NEDD1</sup> is not a general γ-TuRC targeting factor in class IV da neurons, similar to dividing cells where Grip71<sup>NEDD1</sup> is essential for γ-TuRC recruitment to spindle microtubules but has only a minor role in recruitment to centrosomes (Reschen et al., 2012; Vérollet et al., 2006; Zhu et al., 2023). Second, it is consistent with Grip7<sup>NEDD1</sup> being required for Augmin-mediated recruitment of γ-TuRCs in da neurons. This is interesting because the involvement of Grip7<sup>NEDD1</sup> in Augmin-mediated nucleation may influence the angle of the nucleation event. Depletion of NEDD1 from *Arabidopsis* results in a selective loss of branched nucleation events in cortical arrays, with the remaining events being parallel i.e. having a 0° angle. The possibility that Grip7<sup>NEDD1</sup> promotes angled nucleation events fits with an observed average angle of ~36° in *Drosophila* S2 cells (Verma and Maresca, 2019), where Grip7<sup>NEDD1</sup> is required for proper spindle assembly (Dobbelaere et al., 2008; Moutinho-Pereira et al., 2013; Vérollet et al., 2006). Whether Grip7<sup>NEDD1</sup> promotes branched nucleation events in class IV neurons remains to be determined, but such events would allow daughter microtubules to grow directly into a nascent dendrite when nucleated at the branch point (Figure 6A). Branched nucleation events occurring elsewhere may be accommodated if the daughter microtubule can bend within the constraints of the cell membrane. In cultured
446

branchpoint could be stochastic, helping to explain why many nascent dendrites retract and disappear, or it could be influenced by the presence of other regulators of branching, such as Arp2/3 (Stürner et al., 2019); 2) Augmin could amplify microtubules throughout the arbor, increasing the possibility that microtubules turn into nascent dendrites (Figure 6B), although our EB1-GFP data shows that microtubule growth is actually increased in secondary and tertiary dendrites after Augmin depletion; 3) Augmin may amplify anterograde microtubules already present within nascent dendrites (Figure 6C). An alternative possibility is that Augmin helps maintain correct microtubule polarity to ensure the proper delivery of material needed for new branching events (Figure 6D). Future studies will be required to elucidate the exact molecular mechanism underlying Augmin's role in dendrite branching.

447

448

449

450

451

452

453

454

455

456

457

458

459

460

461

462

463

464

465

466

467

468

469

470

471

472

473

474

475

476

477

Intriguingly, we find that Grip7¹<sup>NEDD1</sup> depletion results in similar phenotypes to Augmin depletion. However, both these phenotypes are weaker than those resulting from γ-TuRC depletion. Firstly, this suggests that the γ-TuRC protein Grip71<sup>NEDD1</sup> is not a general γ-TuRC targeting factor in class IV da neurons, similar to dividing cells where Grip71<sup>NEDD1</sup> is essential for γ-TuRC recruitment to spindle microtubules but has only a minor role in recruitment to centrosomes (Reschen et al., 2012; Vérollet et al., 2006; Zhu et al., 2023). Second, it is consistent with Grip7<sup>NEDD1</sup> being required for Augmin-mediated recruitment of γ-TuRCs in da neurons. This is interesting because the involvement of Grip7<sup>NEDD1</sup> in Augmin-mediated nucleation may influence the angle of the nucleation event. Depletion of NEDD1 from *Arabidopsis* results in a selective loss of branched nucleation events in cortical arrays, with the remaining events being parallel i.e. having a 0° angle. The possibility that Grip7<sup>NEDD1</sup> promotes angled nucleation events fits with an observed average angle of ~36° in *Drosophila* S2 cells (Verma and Maresca, 2019), where Grip7<sup>NEDD1</sup> is required for proper spindle assembly (Dobbelaere et al., 2008; Moutinho-Pereira et al., 2013; Vérollet et al., 2006). Whether Grip7<sup>NEDD1</sup> promotes branched nucleation events in class IV neurons remains to be determined, but such events would allow daughter microtubules to grow directly into a nascent dendrite when nucleated at the branch point (Figure 6A). Branched nucleation events occurring elsewhere may be accommodated if the daughter microtubule can bend within the constraints of the cell membrane. In cultured
478

mammalian neurons and hippocampal tissue, however, NEDD1 is downregulated at
later developmental stages, despite there still being an interaction between Augmin
and γ-TuRCs (Sánchez-Huertas et al., 2016). Thus, the finer details of Augmin-
mediated nucleation may differ between species.

479

480

481

482

483

An important question is how γ-TuRCs are employed to help regulate different aspects
of class IV dendritic arbor development. While γ-TuRCs appear to promote higher-
order branching via the Augmin pathway, the difference in phenotypes between
grip91<sup>GCP3</sup>-RNAi and Dgt4-RNAi suggest that γ-TuRCs function independently of
Augmin to mediate the growth of primary and secondary dendrites and major
branching events. Perhaps the γ-TuRCs associated with the few proximal Golgi
outposts within primary branchpoints nucleate microtubules that contribute to these
processes. Perhaps a putative MTOC observed close to the tip of extending primary
dendrites (Yoong et al., 2020) recruits γ-TuRCs that ultimately promote the growth
and branching of primary dendrites. Nevertheless, we cannot completely rule out that
Augmin is required for these major growth and branching events and that residual
Augmin left over after RNAi depletion is sufficient to mask any major defects.

484

485

486

487

488

489

490

491

492

493

494

495

496

Our conclusions on the requirements for γ-TuRCs and Augmin were drawn in part
from the analysis of dendritic arbor re-growth in pupae. Here, we found the phenotypes
induced by RNAi expression to be stronger than those observed in larvae. This may
be due to ineffective RNAi during early arbor development in embryos and larvae,
possibly due to persistent maternal contributions of protein and protein complexes.
Indeed, in our hands, γ-tubulin23c null mutant flies can survive until late larval/early
pupal stages, which is presumably only possible due to a persistent maternal
contribution. Thus, future studies may consider analysing phenotypes in pupae as well
as embryos or larvae. For example, examining EB1-GFP comets in pupal neurons
depleted for γ-TuRCs would be informative, as previous studies in larvae report
relatively minor changes, or no changes, in EB1-GFP comet frequency after γ-tubulin
depletion (Nguyen et al., 2014; Ori-McKenney et al., 2012). Moreover, one may see
stronger effects on microtubule dynamics after Augmin depletion. The use of other

497

498

499

500

501

502

503

504

505

506

507

508
478

mammalian neurons and hippocampal tissue, however, NEDD1 is downregulated at
later developmental stages, despite there still being an interaction between Augmin
and γ-TuRCs (Sánchez-Huertas et al., 2016). Thus, the finer details of Augmin-
mediated nucleation may differ between species.

479

480

481

482

483

An important question is how γ-TuRCs are employed to help regulate different aspects
of class IV dendritic arbor development. While γ-TuRCs appear to promote higher-
order branching via the Augmin pathway, the difference in phenotypes between
grip91<sup>GCP3</sup>-RNAi and Dgt4-RNAi suggest that γ-TuRCs function independently of
Augmin to mediate the growth of primary and secondary dendrites and major
branching events. Perhaps the γ-TuRCs associated with the few proximal Golgi
outposts within primary branchpoints nucleate microtubules that contribute to these
processes. Perhaps a putative MTOC observed close to the tip of extending primary
dendrites (Yoong et al., 2020) recruits γ-TuRCs that ultimately promote the growth
and branching of primary dendrites. Nevertheless, we cannot completely rule out that
Augmin is required for these major growth and branching events and that residual
Augmin left over after RNAi depletion is sufficient to mask any major defects.

484

485

486

487

488

489

490

491

492

493

494

495

496

Our conclusions on the requirements for γ-TuRCs and Augmin were drawn in part
from the analysis of dendritic arbor re-growth in pupae. Here, we found the phenotypes
induced by RNAi expression to be stronger than those observed in larvae. This may
be due to ineffective RNAi during early arbor development in embryos and larvae,
possibly due to persistent maternal contributions of protein and protein complexes.
Indeed, in our hands, γ-tubulin23c null mutant flies can survive until late larval/early
pupal stages, which is presumably only possible due to a persistent maternal
contribution. Thus, future studies may consider analysing phenotypes in pupae as well
as embryos or larvae. For example, examining EB1-GFP comets in pupal neurons
depleted for γ-TuRCs would be informative, as previous studies in larvae report
relatively minor changes, or no changes, in EB1-GFP comet frequency after γ-tubulin
depletion (Nguyen et al., 2014; Ori-McKenney et al., 2012). Moreover, one may see
stronger effects on microtubule dynamics after Augmin depletion. The use of other

497

498

499

500

501

502

503

504

505

506

507

508
509

techniques to directly and quickly deplete or perturb proteins within neurons would
also be of value, as was recently done for Kinesin-1 (Xu et al., 2023).

510

511

512

While the importance of γ-TuRCs for the development of class IV da neuron arbors is
now clear, γ-TuRC-independent microtubule nucleation may also play a role. As
mentioned above, most Golgi outposts do not associate with detectable levels of γ-
tubulin, yet EB1-GFP comets do grow out from them (Ori-McKenney et al., 2012; Zhou
et al., 2014). Moreover, microtubule polarity is perturbed when Golgi outposts are mis-
localised in axons even when γ-TuRC proteins have been depleted (Yang and
Wildonger, 2020). It remains to be explored whether this mode of microtubule
nucleation plays an important role in arbor development. Indeed, more studies are
now required to fully elucidate the intricacies of how γ-TuRCs, Augmin and γ-TuRC-
independent microtubule nucleation events are regulated to ensure correct dendritic
arbor growth and branching.

513

514

515

516

517

518

519

520

521

522
509

techniques to directly and quickly deplete or perturb proteins within neurons would
also be of value, as was recently done for Kinesin-1 (Xu et al., 2023).

510

511

512

While the importance of γ-TuRCs for the development of class IV da neuron arbors is
now clear, γ-TuRC-independent microtubule nucleation may also play a role. As
mentioned above, most Golgi outposts do not associate with detectable levels of γ-
tubulin, yet EB1-GFP comets do grow out from them (Ori-McKenney et al., 2012; Zhou
et al., 2014). Moreover, microtubule polarity is perturbed when Golgi outposts are mis-
localised in axons even when γ-TuRC proteins have been depleted (Yang and
Wildonger, 2020). It remains to be explored whether this mode of microtubule
nucleation plays an important role in arbor development. Indeed, more studies are
now required to fully elucidate the intricacies of how γ-TuRCs, Augmin and γ-TuRC-
independent microtubule nucleation events are regulated to ensure correct dendritic
arbor growth and branching.

513

514

515

516

517

518

519

520

521

522
523

524 **Materials and Methods**

525 **Contact for Reagent and Resource Sharing**

526 Further information and requests for resources and reagents should be directed to and
527 will be fulfilled by the Lead Contact, Paul Conduit (paul.conduit@ijm.fr).

528

529 **Experimental Model and Subject Details**

530 All *Drosophila* stocks and crosses were maintained at 18 or 25°C on Iberian fly food
531 made from dry active yeast, agar, and organic pasta flour, supplemented with nipagin,
532 propionic acid, pen/strep and food colouring.

533

534 **Methods**

535 *Drosophila melanogaster* stocks

536 The following fluorescent alleles were used in this study: ppk-CD4-tdGFP (BL 35842),
537 γ-tubulin23C-sfGFP (Tovey et al., 2018), γ-tubulin23c-eGFP (Mukherjee et al., 2020),
538 UAS-EB1-GFP (BL 35512), UAS-mCD8-RFP (gift from Peter Lawrence and Jose
539 Casal). The following Gal4 and RNAi lines were used in this study: IT.Gal4-dgt5
540 (BL77584), ppk-Gal4 (BL 32078), 221-Gal4 (BL 26259), UAS-IVS-myr-GFP (gift from
541 Matthias Landgraf), UAS-grip91<sup>GCP3</sup>-RNAi1 (VDRC 104667), UAS-grip91<sup>GCP3</sup>-RNAi2
542 (BL 31201), UAS-grip84<sup>GCP2</sup>-RNAi1 (VDRC 105640), UAS-grip84<sup>GCP2</sup>-RNAi2 (BL
543 33548), γ-tubulin23C-RNAi1 (BL 31204), γ-tubulin23C-RNAi2 (VDRC 19130), UAS-
544 Dicer 2 (BL 24646), UAS- grip7<sup>NEDD1</sup>-RNAi1 (VDRC v31228), UAS- grip7<sup>NEDD1</sup>-RNAi2
545 (NIG 10346R-1), UAS-dgt2-RNAi1 (BL31729), UAS-dgt2-RNAi2 (v330504), UAS-
546 dgt3-RNAi1 (VDRC 103980), UAS-dgt3-RNAi2 (BL58137), UAS-dgt4-RNAi (VDRC
547 108969), UAS-dgt5-RNAi (BL60366). UAS-γ-tubulin37C-RNAi (BL32513) was used
548 as the control in all cases. We confirmed by PCR that any RNAi line from the KK library
549 did not contain a known aberrant insertion at genomic position 40D (data not shown –
550 this information is now available on the VDRC website anyway), which upregulates
551 the Hippo pathway and is mistakenly present in some KK library constructs (see
552 information on VDRC website).

553
523

524

**Materials and Methods**

525

**Contact for Reagent and Resource Sharing**

526

527

Further information and requests for resources and reagents should be directed to and will be fulfilled by the Lead Contact, Paul Conduit (paul.conduit@ijm.fr).

528

529

**Experimental Model and Subject Details**

530

531

532

All *Drosophila* stocks and crosses were maintained at 18 or 25°C on Iberian fly food made from dry active yeast, agar, and organic pasta flour, supplemented with nipagin, propionic acid, pen/strep and food colouring.

533

534

**Methods**

535

*Drosophila melanogaster* stocks

536

537

538

539

540

541

542

543

544

545

546

547

548

549

550

551

552

The following fluorescent alleles were used in this study: ppk-CD4-tdGFP (BL 35842), γ-tubulin23C-sfGFP (Tovey et al., 2018), γ-tubulin23c-eGFP (Mukherjee et al., 2020), UAS-EB1-GFP (BL 35512), UAS-mCD8-RFP (gift from Peter Lawrence and Jose Casal). The following Gal4 and RNAi lines were used in this study: IT.Gal4-dgt5 (BL77584), ppk-Gal4 (BL 32078), 221-Gal4 (BL 26259), UAS-IVS-myr-GFP (gift from Matthias Landgraf), UAS-grip91<sup>GCP3</sup>-RNAi1 (VDRC 104667), UAS-grip 91<sup>GCP3</sup>-RNAi2 (BL 31201), UAS-grip84<sup>GCP2</sup>-RNAi1 (VDRC 105640), UAS-grip84<sup>GCP2</sup>-RNAi2 (BL 33548), γ-tubulin23C-RNAi1 (BL 31204), γ-tubulin23C-RNAi2 (VDRC 19130), UAS-Dicer 2 (BL 24646), UAS-grip7<sup>NEDD1</sup>-RNAi1 (VDRC v31228), UAS-grip7<sup>NEDD1</sup>-RNAi2 (NIG 10346R-1), UAS-dgt2-RNAi1 (BL31729), UAS-dgt2-RNAi2 (v330504), UAS-dgt3-RNAi1 (VDRC 103980), UAS-dgt3-RNAi2 (BL58137), UAS-dgt4-RNAi (VDRC 108969), UAS-dgt5-RNAi (BL60366). UAS-γ-tubulin37C-RNAi (BL32513) was used as the control in all cases. We confirmed by PCR that any RNAi line from the KK library did not contain a known aberrant insertion at genomic position 40D (data not shown – this information is now available on the VDRC website anyway), which upregulates the Hippo pathway and is mistakenly present in some KK library constructs (see information on VDRC website).

553
For examining the morphology of pupal or larval class IV neurons we used flies
554
expressing one copy of ppk-CD4-tdGFP and containing one copy of UAS-Dicer2 and
555
one copy of the appropriate UAS-driven RNAi line expressed under the control of one
556
copy of ppk-Gal4. For examining the morphology of larval class I neurons we used
557
flies containing one copy of UAS-mCD8-RFP, one copy of UAS-Dicer2, and one copy
558
of the appropriate UAS-driven RNAi line expressed under the control of one copy of
559
221-Gal4. For examining the endogenous localisation of γ-tubulin23C we used flies
560
expressing γ-tubulin23C-sfGFP and γ-tubulin23C-eGFP i.e. 2 copies of γ-tubulin23C-
561
GFP. For examining the localisation of γ-tubulin23C during Grip91<sup>GCP3</sup> depletion we
562
used flies expressing two copies of γ-tubulin23C-(sf/e)GFP (as above) in combination
563
with one copy of UAS-grip91<sup>GCP3</sup> RNAi and UAS-Dicer2 expressed under the control
564
of one copy of 221-Gal4. For examining microtubule dynamics in control-RNAi or dgt4-
565
RNAi neurons we used flies containing one copy of UAS-EB1-GFP, one copy of UAS-
566
Dicer 2 and one copy of either UAS-γ-tubulin37C-RNAi or UAS-dgt4-RNAi expressed
567
under the control of one copy of ppk-Gal4. For examining dgt-5 expression, we used
568
flies containing one copy of IT.Gal4-dgt5 and one copy of UAS-IVS-myr-GFP.
569

570

571 Antibodies

The following primary antibodies were used: anti-GFP mouse monoclonal at 1:250
572
(Roche, 11814460001), Alexa-647 conjugated anti-HRP goat polyclonal at 1:500
573
(Jackson, AB_2338967), and anti-GM130 rabbit polyclonal at 1:300 (Abcam,
574
ab32337). The following secondary antibodies were used: Alexa-488 goat anti-Mouse
575
at 1:500 (ThermoFisher, 10138324), Alexa-568 anti-Rabbit at 1:500 (ThermoFisher,
576
10463022).
577

578

579 Immunostaining and Fixed cell imaging

Dissected larvae were processed as previously described (Mukherjee et al., 2020).
580
Briefly, fillet preparations were fixed in freshly prepared 4% formaldehyde for 20
581
minutes at room temperature and were then washed four times for 10 minutes in PBST
582
(PBS + 0.1% TritonX-100). Blocking was carried out in PBST + 5% BSA for 1 hour at
583
room temperature. Preparations were incubated with appropriate primary antibodies
584
diluted in PBST overnight at 4°C. After washing in PBST for ~8 hours, changing
585
For examining the morphology of pupal or larval class IV neurons we used flies expressing one copy of ppk-CD4-tdGFP and containing one copy of UAS-Dicer2 and one copy of the appropriate UAS-driven RNAi line expressed under the control of one copy of ppk-Gal4. For examining the morphology of larval class I neurons we used flies containing one copy of UAS-mCD8-RFP, one copy of UAS-Dicer2, and one copy of the appropriate UAS-driven RNAi line expressed under the control of one copy of 221-Gal4. For examining the endogenous localisation of γ-tubulin23C we used flies expressing γ-tubulin23C-sfGFP and γ-tubulin23C-eGFP i.e. 2 copies of γ-tubulin23C-GFP. For examining the localisation of γ-tubulin23C during Grip91<sup>GCP3</sup> depletion we used flies expressing two copies of γ-tubulin23C-(sf/e)GFP (as above) in combination with one copy of UAS-grip91<sup>GCP3</sup> RNAi and UAS-Dicer2 expressed under the control of one copy of 221-Gal4. For examining microtubule dynamics in control-RNAi or dgt4-RNAi neurons we used flies containing one copy of UAS-EB1-GFP, one copy of UAS-Dicer 2 and one copy of either UAS-γ-tubulin37C-RNAi or UAS-dgt4-RNAi expressed under the control of one copy of ppk-Gal4. For examining dgt-5 expression, we used flies containing one copy of IT.Gal4-dgt5 and one copy of UAS-IVS-myr-GFP.

570

## Antibodies

The following primary antibodies were used: anti-GFP mouse monoclonal at 1:250 (Roche, 11814460001), Alexa-647 conjugated anti-HRP goat polyclonal at 1:500 (Jackson, AB_2338967), and anti-GM130 rabbit polyclonal at 1:300 (Abcam, ab32337). The following secondary antibodies were used: Alexa-488 goat anti-Mouse at 1:500 (ThermoFisher, 10138324), Alexa-568 anti-Rabbit at 1:500 (ThermoFisher, 10463022).

578

## Immunostaining and Fixed cell imaging

Dissected larvae were processed as previously described (Mukherjee et al., 2020). Briefly, fillet preparations were fixed in freshly prepared 4% formaldehyde for 20 minutes at room temperature and were then washed four times for 10 minutes in PBST (PBS + 0.1% TritonX-100). Blocking was carried out in PBST + 5% BSA for 1 hour at room temperature. Preparations were incubated with appropriate primary antibodies diluted in PBST overnight at 4°C. After washing in PBST for ~8 hours, changing
washes every 30-45 minutes, samples were incubated in secondary antibodies diluted
in PBST overnight at 4°C. The fillet preparations were then washed for ~8 hours,
changing washes every 30-45 minutes in PBST before mounting in Moviol. They were
stored at -20°C and imaged within a week. Neurons within segments A2 to A6 were
imaged. Imaging of γ-tubulin-GFP within the soma was carried out using an Olympus
FV3000 scanning inverted confocal system run by FV-OSR software with a 60X 1.4NA
silicone immersion lens (UPLSAPO60xSilicone). Images of neuronal somas within
IT.Gal4-dgt5 > UAS-IVS-myr-GFP flies were collected on a Zeiss Axio Observer.Z1
inverted CSU-X1 Yokogowa spinning disk system with 2 ORCA Fusion camera
(Hamamatsu) run by Zeiss Zen2 acquisition software using a 60X 1.4NA oil immersion
lens (Zeiss). Z-stacks with 0.5µm spacing were acquired to cover all neuronal soma.

597

598 Live imaging

All imaging was carried out at ambient room temperature (~21°C). Imaging of overall neuronal morphology was carried out on a Leica SP5 point scanning upright confocal system run by LAS AF software with a 20X 0.7NA multi-immersion lens (11506191) with glycerol as the immersion medium. Z-stacks with 1µm spacing were acquired to cover the whole arbor. For imaging class IV v'ada neurons within pupae, white pre-pupae (0h APF) were collected and placed in a sealed petri dish with some moistened tissue paper and aged for the required duration in a 25°C incubator. The staged pupae were stuck on double-sided sticky tape and removed from their pupal casing. The pupae were mounted on a glass slide with a small amount of halocarbon oil and a coverglass was placed gently on top of the pupae using dental wax feet as spacers. The mounted pupae were imaged immediately. For larval neuron morphology, newly hatched (within 1 hour) larvae were collected and placed on a new plate to be staged appropriately at 25°C. The larvae were then placed in a drop of glycerol and gently flattened between a slide and a 22X22mm coverslip, held in place by tape, and imaged immediately. Imaging of EB1-GFP comets and the dynamic morphology of terminal dendrites was carried out on a Leica DMi8 inverted wide-field microscope controlled by µManager software and coupled to a BSI Prime Express monochrome camera (QImaging) and a CoolLED pE-300 Ultra light source using a 63X 1.3NA oil objective (Leica 11506384). Wandering third instar larvae (between ~96h and ~120h AEL) were

600

601

602

603

604

605

606

607

608

609

610

611

612

613

614

615

616

617
washes every 30-45 minutes, samples were incubated in secondary antibodies diluted
in PBST overnight at 4°C. The fillet preparations were then washed for ~8 hours,
changing washes every 30-45 minutes in PBST before mounting in Moviol. They were
stored at -20°C and imaged within a week. Neurons within segments A2 to A6 were
imaged. Imaging of γ-tubulin-GFP within the soma was carried out using an Olympus
FV3000 scanning inverted confocal system run by FV-OSR software with a 60X 1.4NA
silicone immersion lens (UPLSAPO60xSilicone). Images of neuronal somas within
IT.Gal4-dgt5 > UAS-IVS-myr-GFP flies were collected on a Zeiss Axio Observer.Z1
inverted CSU-X1 Yokogowa spinning disk system with 2 ORCA Fusion camera
(Hamamatsu) run by Zeiss Zen2 acquisition software using a 60X 1.4NA oil immersion
lens (Zeiss). Z-stacks with 0.5µm spacing were acquired to cover all neuronal soma.

597

598 Live imaging

All imaging was carried out at ambient room temperature (~21°C). Imaging of overall neuronal morphology was carried out on a Leica SP5 point scanning upright confocal system run by LAS AF software with a 20X 0.7NA multi-immersion lens (11506191) with glycerol as the immersion medium. Z-stacks with 1µm spacing were acquired to cover the whole arbor. For imaging class IV v'ada neurons within pupae, white pre-pupae (0h APF) were collected and placed in a sealed petri dish with some moistened tissue paper and aged for the required duration in a 25°C incubator. The staged pupae were stuck on double-sided sticky tape and removed from their pupal casing. The pupae were mounted on a glass slide with a small amount of halocarbon oil and a coverglass was placed gently on top of the pupae using dental wax feet as spacers. The mounted pupae were imaged immediately. For larval neuron morphology, newly hatched (within 1 hour) larvae were collected and placed on a new plate to be staged appropriately at 25°C. The larvae were then placed in a drop of glycerol and gently flattened between a slide and a 22X22mm coverslip, held in place by tape, and imaged immediately. Imaging of EB1-GFP comets and the dynamic morphology of terminal dendrites was carried out on a Leica DMi8 inverted wide-field microscope controlled by µManager software and coupled to a BSI Prime Express monochrome camera (QImaging) and a CoolLED pE-300 Ultra light source using a 63X 1.3NA oil objective (Leica 11506384). Wandering third instar larvae (between ~96h and ~120h AEL) were

600

601

602

603

604

605

606

607

608

609

610

611

612

613

614

615

616

617
618

placed in a drop of glycerol and gently held between a slide and a 22X22mm coverslip using tape to hold down the coverslip, and imaged immediately. Care was taken not to flatten the larvae too much, as we found that fully squishing the larvae resulted in abnormal neuronal morphology defects. For EB1-GFP, 5 Z-stacks of 0.5µm spacing were acquired every 5 seconds. For analysing the dynamic morphology of terminal dendrites, 10 Z-stacks of 1µm spacing were acquired every 5 minutes for a total of 30 minutes.

619

620

621

622

623

624

625

626

## Image analysis and statistics

627

All images were processed in Fiji (ImageJ). Images of neuronal morphology, γ-tubulin-GFP within soma, the dgt-5 Gal4 trap, and the dynamic morphology of terminal dendrites are maximum intensity projections of Z stacks. To determine peripheral area for a neuron, a line was manually drawn around the neuronal field by joining the tips of dendrites at the edge of the arbor. In larvae, care was taken not to draw the line within large empty regions between dendrites in order to better measure the outgrowth of primary and secondary dendrites. Dendrite tips were counted manually for each neuron. To measure the length of the longest primary dendrite in pupae, a line was manually drawn along the dendrite and its length measured in Fiji. To measure the total length of the primary and secondary dendrites in pupae, segmented lines were manually drawn along the dendrites and their lengths measured in Fiji and then summed. Amira software was used to measure the total arbor length and tip number of class I da neurons.

628

629

630

631

632

633

634

635

636

637

638

639

640

641

For the analysis of the γ-tubulin-GFP signal at somatic Golgi stacks in control versus grip91<sup>GCP3</sup>-RNAi neurons, ROIs were drawn around individual Golgi stacks and their sum fluorescence intensities were background corrected using a mean cytosolic intensity measurement. EB1 comets were manually scored for their number and direction within different types of dendrites. Data was processed in Microsoft Excel. Statistical analysis and graph production were performed using GraphPad Prism. For analysing the expression of IT.Gal4-dgt5 > UAS-IVS-myr-GFP, intensity measurements within the soma were used to determine whether the expression was similar, higher, or lower between class I and class IV da neurons.

642

643

644

645

646

647

648

649
618

placed in a drop of glycerol and gently held between a slide and a 22X22mm coverslip using tape to hold down the coverslip, and imaged immediately. Care was taken not to flatten the larvae too much, as we found that fully squishing the larvae resulted in abnormal neuronal morphology defects. For EB1-GFP, 5 Z-stacks of 0.5µm spacing were acquired every 5 seconds. For analysing the dynamic morphology of terminal dendrites, 10 Z-stacks of 1µm spacing were acquired every 5 minutes for a total of 30 minutes.

619

620

621

622

623

624

625

626

## Image analysis and statistics

627

All images were processed in Fiji (ImageJ). Images of neuronal morphology, γ-tubulin-GFP within soma, the dgt-5 Gal4 trap, and the dynamic morphology of terminal dendrites are maximum intensity projections of Z stacks. To determine peripheral area for a neuron, a line was manually drawn around the neuronal field by joining the tips of dendrites at the edge of the arbor. In larvae, care was taken not to draw the line within large empty regions between dendrites in order to better measure the outgrowth of primary and secondary dendrites. Dendrite tips were counted manually for each neuron. To measure the length of the longest primary dendrite in pupae, a line was manually drawn along the dendrite and its length measured in Fiji. To measure the total length of the primary and secondary dendrites in pupae, segmented lines were manually drawn along the dendrites and their lengths measured in Fiji and then summed. Amira software was used to measure the total arbor length and tip number of class I da neurons.

628

629

630

631

632

633

634

635

636

637

638

639

640

641

For the analysis of the γ-tubulin-GFP signal at somatic Golgi stacks in control versus grip91<sup>GCP3</sup>-RNAi neurons, ROIs were drawn around individual Golgi stacks and their sum fluorescence intensities were background corrected using a mean cytosolic intensity measurement. EB1 comets were manually scored for their number and direction within different types of dendrites. Data was processed in Microsoft Excel. Statistical analysis and graph production were performed using GraphPad Prism. For analysing the expression of IT.Gal4-dgt5 > UAS-IVS-myr-GFP, intensity measurements within the soma were used to determine whether the expression was similar, higher, or lower between class I and class IV da neurons.

642

643

644

645

646

647

648

649
650

651

652

653

654

655

656

657

658

659

660

661

662

663

664

665

666

667

668

669

When parametric tests were used, tests for normality were first performed using D'agostino and Pearson tests, Anderson-Darling tests, Shapiro-Wilk tests, and Kolmogorov-Smirnov tests. Datasets were considered to be normally distributed when at least one test passed the normality test. All T-tests were 2-sided. When using One-way ANOVA we assumed equal SDs. We corrected for multiple comparisons using Šídák hypothesis testing and multiplicity corrected p values for each comparison were reported. When using Kruskal-Wallis tests we corrected for multiple comparisons using Dunn's hypothesis testing and multiplicity corrected p values for each comparison were reported. The following tests were performed to make comparisons between datasets: For Figures 1C,D and 3B-D we used a one-way ANOVA with correction for multiple comparisons. For Figures 1E,F,H, 2B,D, 4B,D,E, 5B,D, we used we used t-tests. For Figures 1G we used a Kolmogorov-Smirnov test. For Figure 3F we used a Kruskal-Wallis test. For Figure 4F we used a Chi-squared test. For Figures 3D,E,G we use One-way ANOVA. For Figure 4G,J we used Fisher's exact tests. For Figure 4I we used a Mann-Whitney test. The data in Figure S3G was determined to be log-normally distributed and so the data was first log transformed by taking the log₁₀ values before performing a t-test.

670

671

672

673

674

675

676

677

678

679

680

681

RT-qPCR

For each of 3 biological replicates, wing imaginal discs were dissected from 25 males flies then washed with Phosphatate Buffer Saline (PBS) Buffer and stored at -70°C in Trizol (Thermofischer) until use. Total RNA was isolated from the sample using RNeasy Mini Kit (Qiagen). cDNA was synthesized using Superscript III Reverse Transcription Kit (Invitrogen). cDNA synthesis for each biological replicate was repeated 3 times on 3 separate days to generate 9 samples for quantitative RT-PCR. Quantitative RT-PCR was run in triplicate for each of the 9 samples in 15 µL reactions in a real-time PCR system (LightCycler® 480 Multiwell Plate 384, Roche, Life Science) using SYBR-Green qPCR mix (Roche, Life Science). qPCR software was used for analyzing cycle threshold (Ct) values. Fold change in mRNA levels (expressed as 2⁻⁰⁻⁰⁾ was normalized to the mean expression of 4 housekeeping genes: α-tubulin84b, Ribosomal Protein S2 (SOP), Ribosomal Protein L32 (RPL32) and Glyceraldehyde 3-
650

651

652

653

654

655

656

657

658

659

660

661

662

663

664

665

666

667

668

669

When parametric tests were used, tests for normality were first performed using D'agostino and Pearson tests, Anderson-Darling tests, Shapiro-Wilk tests, and Kolmogorov-Smirnov tests. Datasets were considered to be normally distributed when at least one test passed the normality test. All T-tests were 2-sided. When using One-way ANOVA we assumed equal SDs. We corrected for multiple comparisons using Šídák hypothesis testing and multiplicity corrected p values for each comparison were reported. When using Kruskal-Wallis tests we corrected for multiple comparisons using Dunn's hypothesis testing and multiplicity corrected p values for each comparison were reported. The following tests were performed to make comparisons between datasets: For Figures 1C,D and 3B-D we used a one-way ANOVA with correction for multiple comparisons. For Figures 1E,F,H, 2B,D, 4B,D,E, 5B,D, we used we used t-tests. For Figures 1G we used a Kolmogorov-Smirnov test. For Figure 3F we used a Kruskal-Wallis test. For Figure 4F we used a Chi-squared test. For Figures 3D,E,G we use One-way ANOVA. For Figure 4G,J we used Fisher's exact tests. For Figure 4I we used a Mann-Whitney test. The data in Figure S3G was determined to be log-normally distributed and so the data was first log transformed by taking the log₁₀ values before performing a t-test.

670

671

672

673

674

675

676

677

678

679

680

681

RT-qPCR

For each of 3 biological replicates, wing imaginal discs were dissected from 25 males flies then washed with Phosphatate Buffer Saline (PBS) Buffer and stored at -70°C in Trizol (Thermofischer) until use. Total RNA was isolated from the sample using RNeasy Mini Kit (Qiagen). cDNA was synthesized using Superscript III Reverse Transcription Kit (Invitrogen). cDNA synthesis for each biological replicate was repeated 3 times on 3 separate days to generate 9 samples for quantitative RT-PCR. Quantitative RT-PCR was run in triplicate for each of the 9 samples in 15 µL reactions in a real-time PCR system (LightCycler® 480 Multiwell Plate 384, Roche, Life Science) using SYBR-Green qPCR mix (Roche, Life Science). qPCR software was used for analyzing cycle threshold (Ct) values. Fold change in mRNA levels (expressed as 2⁻⁰⁻⁰⁾ was normalized to the mean expression of 4 housekeeping genes: α-tubulin84b, Ribosomal Protein S2 (SOP), Ribosomal Protein L32 (RPL32) and Glyceraldehyde 3-
682 phosphate dehydrogenase (GAPDH-2). The sequence of the primers used is given in

683 Table S3. Source datafile 1 contains all the raw data.
682 phosphate dehydrogenase (GAPDH-2). The sequence of the primers used is given in

683 Table S3. Source datafile 1 contains all the raw data.
684 Acknowledgements

685 This research was supported by the Centre National de la Recherche Scientifique
686 CNRS, Université Paris Cité, a Wellcome Trust and Royal Society Sir Henry Dale
687 fellowship awarded to PTC [105653/Z/14/Z], an Isaac Newton Trust Research grant
688 (18.23(p)) awarded to PTC, by a Chaire d'excellence grant from the IdEx Université
689 Paris Cité ANR-18-IDEX-0001 awarded to PTC, by an ATIP Avenir award funded by
690 the Fondation Bettencourt Schueller awarded to PTC. We thank Melissa Rolls, Bing
691 Ye and Yuh-Nung Jan for sharing fly lines. The work benefited from the Imaging
692 Facility, Department of Zoology, University of Cambridge, supported by Matt Wayland
693 and a Sir Isaac Newton Trust Research Grant (18.07ii(c)). We also acknowledge the
694 ImagoSeine core facility of Institut Jacques Monod, member of France-BioImaging
695 (ANR-10-INBS-04) and IBiSA, with the support of Labex "Who Am I", Inserm Plan
696 Cancer, Region Ile-de-France and Fondation Bettencourt Schueller. For the purpose
697 of Open Access, the author has applied a CC BY public copyright license to any Author
698 Accepted Manuscript version arising from this submission. The authors declare no
699 competing financial interests.

700

701 Data availability statement

702 The Excel sheets and image files containing the raw data underlying all figures are
703 available from the corresponding author upon reasonable request.

704
684 Acknowledgements

685 This research was supported by the Centre National de la Recherche Scientifique
686 CNRS, Université Paris Cité, a Wellcome Trust and Royal Society Sir Henry Dale
687 fellowship awarded to PTC [105653/Z/14/Z], an Isaac Newton Trust Research grant
688 (18.23(p)) awarded to PTC, by a Chaire d'excellence grant from the IdEx Université
689 Paris Cité ANR-18-IDEX-0001 awarded to PTC, by an ATIP Avenir award funded by
690 the Fondation Bettencourt Schueller awarded to PTC. We thank Melissa Rolls, Bing
691 Ye and Yuh-Nung Jan for sharing fly lines. The work benefited from the Imaging
692 Facility, Department of Zoology, University of Cambridge, supported by Matt Wayland
693 and a Sir Isaac Newton Trust Research Grant (18.07ii(c)). We also acknowledge the
694 ImagoSeine core facility of Institut Jacques Monod, member of France-BioImaging
695 (ANR-10-INBS-04) and IBiSA, with the support of Labex "Who Am I", Inserm Plan
696 Cancer, Region Ile-de-France and Fondation Bettencourt Schueller. For the purpose
697 of Open Access, the author has applied a CC BY public copyright license to any Author
698 Accepted Manuscript version arising from this submission. The authors declare no
699 competing financial interests.

700

701 Data availability statement

702 The Excel sheets and image files containing the raw data underlying all figures are
703 available from the corresponding author upon reasonable request.

704
Figure Legends

**Figure 1.** γ-TuRC depletion results in severe dendritic morphology defects in the dendritic arbors of pupal class IV neurons. (A) Confocal images show pupal class IV v'ada neurons expressing CD4-tdGFP at different times APF, as indicated, using ppk-Gal4. Black dotted box indicates a single dendritic field. Note that the neurons are from different animals but were imaged within the same abdominal segment. (B) Confocal images show pupal class IV v'ada neurons at 72h APF expressing CD4-tdGFP, UAS-Dicer2 and different UAS-RNAi lines, as indicated, using ppk-Gal4. (C,D) Graphs show the perimeter area of dendritic arbors (µm²) (C) and the number of dendrite tips (D) of pupal class IV v'ada neurons at 72h APF expressing CD4-tdGFP, UAS-Dicer2 and different UAS-RNAi lines, as indicated, using ppk-Gal4. Each point on the graph represents a different neuron. n=14, 18, 18, 6, 11, 17, 6 for the datasets from left to right on the graph in (C) and n=12, 18, 18, 6, 10, 17, 6 for the datasets from left to right on the graph in (D). Mean and 95% CI's are indicated. One-way ANOVA was used to compare the control dataset to each experimental dataset. P values are indicated above each experimental dataset. Note how most γ-TuRC RNAi constructs lead to a reduction in arbor area and tip number but that expressing grip91<sup>GCP3</sup>-RNAi1 results in the strongest reduction in both indicators. (E,F) Graphs show the lengths of the longest primary dendrites (E) or the combined length of primary and secondary dendrites (F) for control-RNAi (n=14) and grip91<sup>GCP3</sup>-RNAi1 (n=17) neurons. Mean and 95% CI's are indicated. Two-sided t-tests were used to compare the datasets. (G) Graph shows the relative proportion of primary and secondary branch points for control-RNAi (n=13) and grip91<sup>GCP3</sup>-RNAi1 (n=16) neurons. Each segment represents the percentage for a given number of branch points. A two-sample Kolmogorov-Smirnov test was used to compare the datasets. (H) Graph shows the ratio between primary/secondary dendrite length and branch point number for control-RNAi (n=13) and grip91<sup>GCP3</sup>-RNAi1 (n=16) neurons. Mean and 95% CI's are indicated. A two-sided t-test was used to compare the datasets.

**Figure 2.** γ-TuRC depletion affects higher-order branching at 72h AEL but both primary and secondary dendrite growth and higher-order branching at 96h AEL. (A-D) Confocal images (A,C) show larval class IV ddaC neurons at either 72h AEL (A)
Figure Legends

**Figure 1.** γ-TuRC depletion results in severe dendritic morphology defects in the dendritic arbors of pupal class IV neurons. (A) Confocal images show pupal class IV v'ada neurons expressing CD4-tdGFP at different times APF, as indicated, using ppk-Gal4. Black dotted box indicates a single dendritic field. Note that the neurons are from different animals but were imaged within the same abdominal segment. (B) Confocal images show pupal class IV v'ada neurons at 72h APF expressing CD4-tdGFP, UAS-Dicer2 and different UAS-RNAi lines, as indicated, using ppk-Gal4. (C,D) Graphs show the perimeter area of dendritic arbors (μm²) (C) and the number of dendrite tips (D) of pupal class IV v'ada neurons at 72h APF expressing CD4-tdGFP, UAS-Dicer2 and different UAS-RNAi lines, as indicated, using ppk-Gal4. Each point on the graph represents a different neuron. n=14, 18, 18, 6, 11, 17, 6 for the datasets from left to right on the graph in (C) and n=12, 18, 18, 6, 10, 17, 6 for the datasets from left to right on the graph in (D). Mean and 95% CI's are indicated. One-way ANOVA was used to compare the control dataset to each experimental dataset. P values are indicated above each experimental dataset. Note how most γ-TuRC RNAi constructs lead to a reduction in arbor area and tip number but that expressing grip91<sup>GCP3</sup>-RNAi1 results in the strongest reduction in both indicators. (E,F) Graphs show the lengths of the longest primary dendrites (E) or the combined length of primary and secondary dendrites (F) for control-RNAi (n=14) and grip91<sup>GCP3</sup>-RNAi1 (n=17) neurons. Mean and 95% CI's are indicated. Two-sided t-tests were used to compare the datasets. (G) Graph shows the relative proportion of primary and secondary branch points for control-RNAi (n=13) and grip91<sup>GCP3</sup>-RNAi1 (n=16) neurons. Each segment represents the percentage for a given number of branch points. A two-sample Kolmogorov-Smirnov test was used to compare the datasets. (H) Graph shows the ratio between primary/secondary dendrite length and branch point number for control-RNAi (n=13) and grip91<sup>GCP3</sup>-RNAi1 (n=16) neurons. Mean and 95% CI's are indicated. A two-sided t-test was used to compare the datasets.

**Figure 2.** γ-TuRC depletion affects higher-order branching at 72h AEL but both primary and secondary dendrite growth and higher-order branching at 96h AEL. (A-D) Confocal images (A,C) show larval class IV ddaC neurons at either 72h AEL (A)
or 96h AEL (C) expressing CD4-tdGFP, UAS-Dicer2 and either control-RNAi or grip91<sup>GCP3</sup>-RNAi, as indicated, using ppk-Gal4. Enlargements of insets for control-RNAi (blue) and grip91<sup>GCP3</sup>-RNAi (magenta) are shown on the right of each panel. Orange arrowheads in (C) indicate large gaps between adjacent neurons. Scale bars are indicated within the images. Graphs (B,D) show the perimeter area of dendritic arbors (µm²) (left axes) and the number of dendrite tips (right axes) of larval class IV ddaC neurons at either 72h AEL (B) or 96h AEL (D) expressing CD4-tdGFP, UAS-Dicer2 and either control-RNAi or grip91<sup>GCP3</sup>-RNAi, as indicated, using ppk-Gal4. Each point on the graph represents a different neuron, n=16 and 13 for control-RNAi and grip91<sup>GCP3</sup>-RNAi respectively in (B), n=16, 14, 18, and 14 for the datasets from left to right in the graph in (D). Mean and 95% CI's are indicated. Two-sided t-tests were used to compare the datasets. P values are indicated for each comparison.

749

**Figure 3.** Augmin depletion reduces higher-order branching in the dendritic arbors of pupal class IV neurons. (A) Confocal images show pupal class IV v'ada neurons at 72h APF expressing CD4-tdGFP, UAS-Dicer2 and different UAS-RNAi lines, as indicated, using ppk-Gal4. Scale bars are indicated within the images. (B,C) Graphs show the perimeter area of dendritic arbors (µm²) (B) and the number of dendrite tips (C) of pupal class IV v'ada neurons at 72h APF expressing CD4-tdGFP, UAS-Dicer2 and different UAS-RNAi lines, as indicated, using ppk-Gal4. Each point on the graph represents a different neuron. n=36, 24, 23, 23, 17, 19, 12, and 18 for the datasets from left to right in the graph in (B) and n=33, 19, 6, 22, 22, 17, 16, 12, and 18 for the datasets from left to right in the graph in (C). Mean and 95% CI's are indicated. One-way ANOVA was used to compare the control dataset to each experimental dataset. P values are indicated above each experimental dataset. Note how most Augmin RNAi constructs lead to a reduction in arbor area and tip number but that expressing dgt4-RNAi results in the strongest reduction in both indicators. (D,E) Graphs show the lengths of the longest primary dendrites (D) or the combined length of primary and secondary dendrites (E) for control-RNAi (n=20), grip71<sup>NEDD1</sup>-RNAi1 (n=17) and dgt4-RNAi (n=12) neurons. Mean and 95% CI's are indicated. One-way ANOVA was used to compare the control dataset to each experimental dataset. (F) Graph shows the relative proportion of primary and secondary branch points for
or 96h AEL (C) expressing CD4-tdGFP, UAS-Dicer2 and either control-RNAi or grip91<sup>GCP3</sup>-RNAi, as indicated, using ppk-Gal4. Enlargements of insets for control-RNAi (blue) and grip91<sup>GCP3</sup>-RNAi (magenta) are shown on the right of each panel. Orange arrowheads in (C) indicate large gaps between adjacent neurons. Scale bars are indicated within the images. Graphs (B,D) show the perimeter area of dendritic arbors (µm²) (left axes) and the number of dendrite tips (right axes) of larval class IV ddaC neurons at either 72h AEL (B) or 96h AEL (D) expressing CD4-tdGFP, UAS-Dicer2 and either control-RNAi or grip91<sup>GCP3</sup>-RNAi, as indicated, using ppk-Gal4. Each point on the graph represents a different neuron, n=16 and 13 for control-RNAi and grip91<sup>GCP3</sup>-RNAi respectively in (B), n=16, 14, 18, and 14 for the datasets from left to right in the graph in (D). Mean and 95% CI's are indicated. Two-sided t-tests were used to compare the datasets. P values are indicated for each comparison.

749

Figure 3. Augmin depletion reduces higher-order branching in the dendritic arbors of pupal class IV neurons. (A) Confocal images show pupal class IV v'ada neurons at 72h APF expressing CD4-tdGFP, UAS-Dicer2 and different UAS-RNAi lines, as indicated, using ppk-Gal4. Scale bars are indicated within the images. (B,C) Graphs show the perimeter area of dendritic arbors (µm²) (B) and the number of dendrite tips (C) of pupal class IV v'ada neurons at 72h APF expressing CD4-tdGFP, UAS-Dicer2 and different UAS-RNAi lines, as indicated, using ppk-Gal4. Each point on the graph represents a different neuron. n=36, 24, 23, 23, 17, 19, 12, and 18 for the datasets from left to right in the graph in (B) and n=33, 19, 6, 22, 22, 17, 16, 12, and 18 for the datasets from left to right in the graph in (C). Mean and 95% CI's are indicated. One-way ANOVA was used to compare the control dataset to each experimental dataset. P values are indicated above each experimental dataset. Note how most Augmin RNAi constructs lead to a reduction in arbor area and tip number but that expressing dgt4-RNAi results in the strongest reduction in both indicators. (D,E) Graphs show the lengths of the longest primary dendrites (D) or the combined length of primary and secondary dendrites (E) for control-RNAi (n=20), grip71<sup>NEDD1</sup>-RNAi1 (n=17) and dgt4-RNAi (n=12) neurons. Mean and 95% CI's are indicated. One-way ANOVA was used to compare the control dataset to each experimental dataset. (F) Graph shows the relative proportion of primary and secondary branch points for
769 control-RNAi (n=20), grip7<sup>NEDD1</sup>-RNAi1 (n=17) and dgt4-RNAi (n=12) neurons. Each
770 segment represents the percentage for a given number of branch points. A Kruskal-
771 Wallis test was used to compare each condition to the control. (G) Graph shows the
772 ratio between primary/secondary dendrite length and branch point number for control-
773 RNAi (n=20), grip7<sup>NEDD1</sup>-RNAi1 (n=17) and dgt4-RNAi (n=12) neurons. Mean and 95%
774 CI's are indicated. One-way ANOVA was used to compare the datasets.
775

775

776 **Figure 4. Augmin depletion affects higher-order branching in larval class IV da**
777 **neurons. (A-D)** Confocal images (A,C) show larval class IV ddaC neurons at either
778 96h AEL (A) or 120h AEL (C) expressing CD4-tdGFP, UAS-Dicer2 and either control-
779 RNAi or dgt4-RNAi, as indicated, using ppk-Gal4. Enlargements of insets for control-
780 RNAi (blue) and dgt4-RNAi (yellow) are shown on the right of each panel. Scale bars
781 are indicated within the images. Graphs (B,D) show the perimeter area of dendritic
782 arbors (µm²) (left axes) and the number of dendrite tips (right axes) of larval class IV
783 ddaC neurons at either 96h AEL (B) or 120h AEL (D) expressing CD4-tdGFP, UAS-
784 Dicer2 and either control-RNAi or dgt4-RNAi, as indicated, using ppk-Gal4. Each point
785 on the graph represents a different neuron. n=17, 13, 18, 15 for the datasets from left
to right on the graph (B) and n=13, 14, 13, 15 for the datasets from left to right on the
786 graph (D). Mean and 95% CI's are indicated. Two-sided t-tests were used to compare
787 the datasets. P values are indicated for each comparison. (E) Graph shows the
788 average lengths of terminal dendrites for control and dgt4-RNAi neurons. Each data
789 point represents the average value from an individual neuron. n=10 neurons for both
790 conditions. Mean and 95% CI's are indicated. A two-sided t-test was used to compare
791 the datasets. (F) Graph shows the relative proportion of terminal dendrites of a given
792 length (as indicated below) from control-RNAi (blue) or dgt4-RNAi (purple) neurons.
793 Number of dendrites analysed: 726 for control-RNAi and 536 for dgt4-RNAi. Note the
794 shift in distribution towards shorter dendrites in dgt4-RNAi neurons. (G) Graphs show
795 the proportion of retrograde (white) or anterograde (grey) EB1-GFP comets in either
796 major proximal dendrites, distal secondary dendrites, distal tertiary dendrites, or distal
797 terminal dendrites, as indicated. n=(dendrites;neurons): (216;6), (201;7), (101;5),
798 (208;7), (89;7), (248;7), (45;7), (37;7) for the datasets from left to right on the graph.
799 Fisher's exact tests were used to compare the proportions. P values are indicated
800
769 control-RNAi (n=20), grip7<sup>NEDD1</sup>-RNAi1 (n=17) and dgt4-RNAi (n=12) neurons. Each
770 segment represents the percentage for a given number of branch points. A Kruskal-
771 Wallis test was used to compare each condition to the control. (G) Graph shows the
772 ratio between primary/secondary dendrite length and branch point number for control-
773 RNAi (n=20), grip7<sup>NEDD1</sup>-RNAi1 (n=17) and dgt4-RNAi (n=12) neurons. Mean and 95%
774 CI's are indicated. One-way ANOVA was used to compare the datasets.
775

775

776 **Figure 4. Augmin depletion affects higher-order branching in larval class IV da**
777 **neurons. (A-D)** Confocal images (A,C) show larval class IV ddaC neurons at either
778 96h AEL (A) or 120h AEL (C) expressing CD4-tdGFP, UAS-Dicer2 and either control-
779 RNAi or dgt4-RNAi, as indicated, using ppk-Gal4. Enlargements of insets for control-
780 RNAi (blue) and dgt4-RNAi (yellow) are shown on the right of each panel. Scale bars
781 are indicated within the images. Graphs (B,D) show the perimeter area of dendritic
782 arbors (µm²) (left axes) and the number of dendrite tips (right axes) of larval class IV
783 ddaC neurons at either 96h AEL (B) or 120h AEL (D) expressing CD4-tdGFP, UAS-
784 Dicer2 and either control-RNAi or dgt4-RNAi, as indicated, using ppk-Gal4. Each point
785 on the graph represents a different neuron. n=17, 13, 18, 15 for the datasets from left
to right on the graph (B) and n=13, 14, 13, 15 for the datasets from left to right on the
786 graph (D). Mean and 95% CI's are indicated. Two-sided t-tests were used to compare
787 the datasets. P values are indicated for each comparison. (E) Graph shows the
788 average lengths of terminal dendrites for control and dgt4-RNAi neurons. Each data
789 point represents the average value from an individual neuron. n=10 neurons for both
790 conditions. Mean and 95% CI's are indicated. A two-sided t-test was used to compare
791 the datasets. (F) Graph shows the relative proportion of terminal dendrites of a given
792 length (as indicated below) from control-RNAi (blue) or dgt4-RNAi (purple) neurons.
793 Number of dendrites analysed: 726 for control-RNAi and 536 for dgt4-RNAi. Note the
794 shift in distribution towards shorter dendrites in dgt4-RNAi neurons. (G) Graphs show
795 the proportion of retrograde (white) or anterograde (grey) EB1-GFP comets in either
796 major proximal dendrites, distal secondary dendrites, distal tertiary dendrites, or distal
797 terminal dendrites, as indicated. n=(dendrites;neurons): (216;6), (201;7), (101;5),
798 (208;7), (89;7), (248;7), (45;7), (37;7) for the datasets from left to right on the graph.
799 Fisher's exact tests were used to compare the proportions. P values are indicated
800
above the graphs. (H) Graph shows the number of EB1-GFP comets per 10µm per min in either control-RNAi or dgt4-RNAi distal secondary dendrites, tertiary dendrites, or terminal dendrites. Secondary and tertiary dendrites were grouped together as they displayed similar EB1-GFP comet frequencies. Each point on the graph represents a different neuron. n=9, 11, 9, 12 for the datasets from left to right on the graph. Mean and 95% CI's are indicated. Unpaired t-tests were used to compare control-RNAi and dgt4-RNAi neurons for each dendrite type. P values are indicated above the graph. (I) Graph shows the frequency at which new terminal branches formed within control and dgt4-RNAi neurons during a period of 30 minutes. Each data point represents an individual neuron. n=5 neurons for both conditions. Median and interquartile range are indicated. A two-sided t-test was used to compare the datasets. A two-tailed Mann-Whitney test was used to compare the datasets. P value indicated above the graph. (J) Graph shows the proportion of terminal dendrites that remain (green) or disappear during the 30 mins in either control-RNAi or dgt4-RNAi neurons, as indicated. A Fisher's exact test was used to compare the proportions. P value indicated above the graph.

**Figure 5. Augmin depletion does not affect dendritic branching in larval class I dda neurons. (A-D)** Confocal images (A,C) show larval class I ddaE neurons at either 96h AEL (A) or 120h AEL (C) expressing CD4-tdGFP, UAS-Dicer2 and either control-RNAi or dgt4-RNAi, as indicated, using 221-Gal4. Enlargements of insets for control-RNAi (blue) and dgt4-RNAi (purple) are shown on the right of each panel. Scale bars are indicated within the images. Graphs (B,D) show the total arbor length (µm) (left axes) and the number of dendrite tips (right axes) of larval class I ddaE neurons at either 96h AEL (B) or 120h AEL (D) expressing CD4-tdGFP, UAS-Dicer2 and either control-RNAi or dgt4-RNAi, as indicated, using 221-Gal4. Each point on the graph represents a different neuron. n=15, 16, 15, and 16 for the datasets from left to right on the graph (B) and n=18, 16, 18, and 16 for the datasets from left to right on the graph (D). Mean and 95% CI's are indicated. Two-sided t-tests were used to compare the datasets. P values are indicated for each comparison.
above the graphs. (H) Graph shows the number of EB1-GFP comets per 10µm per min in either control-RNAi or dgt4-RNAi distal secondary dendrites, tertiary dendrites, or terminal dendrites. Secondary and tertiary dendrites were grouped together as they displayed similar EB1-GFP comet frequencies. Each point on the graph represents a different neuron. n=9, 11, 9, 12 for the datasets from left to right on the graph. Mean and 95% CI's are indicated. Unpaired t-tests were used to compare control-RNAi and dgt4-RNAi neurons for each dendrite type. P values are indicated above the graph. (I) Graph shows the frequency at which new terminal branches formed within control and dgt4-RNAi neurons during a period of 30 minutes. Each data point represents an individual neuron. n=5 neurons for both conditions. Median and interquartile range are indicated. A two-sided t-test was used to compare the datasets. A two-tailed Mann-Whitney test was used to compare the datasets. P value indicated above the graph. (J) Graph shows the proportion of terminal dendrites that remain (green) or disappear during the 30 mins in either control-RNAi or dgt4-RNAi neurons, as indicated. A Fisher's exact test was used to compare the proportions. P value indicated above the graph.

**Figure 5. Augmin depletion does not affect dendritic branching in larval class I dda neurons. (A-D)** Confocal images (A,C) show larval class I ddaE neurons at either 96h AEL (A) or 120h AEL (C) expressing CD4-tdGFP, UAS-Dicer2 and either control-RNAi or dgt4-RNAi, as indicated, using 221-Gal4. Enlargements of insets for control-RNAi (blue) and dgt4-RNAi (purple) are shown on the right of each panel. Scale bars are indicated within the images. Graphs (B,D) show the total arbor length (µm) (left axes) and the number of dendrite tips (right axes) of larval class I ddaE neurons at either 96h AEL (B) or 120h AEL (D) expressing CD4-tdGFP, UAS-Dicer2 and either control-RNAi or dgt4-RNAi, as indicated, using 221-Gal4. Each point on the graph represents a different neuron. n=15, 16, 15, and 16 for the datasets from left to right on the graph (B) and n=18, 16, 18, and 16 for the datasets from left to right on the graph (D). Mean and 95% CI's are indicated. Two-sided t-tests were used to compare the datasets. P values are indicated for each comparison.
**Figure 6.** Non-mutually exclusive possibilities for how Augmin may promote **higher-order dendritic branching.** Cartoons representing how Augmin-mediated microtubule nucleation may promote the formation of terminal dendrites from either secondary or tertiary dendrites in the distal region of the class IV da neuron. (A) Augmin-γ-TuRC (purple and pink respectively) may assemble at the base of nascent dendrites and nucleate microtubules at angles that allow the daughter microtubules to grow into and stabilise the nascent dendrite. (B) Augmin-γ-TuRC may be required to amplify microtubules throughout the dendritic arbor, increasing the likelihood that microtubules turn into and stabilise nascent dendrites. (C) Augmin-γ-TuRC may be required to amplify anterograde microtubules within nascent dendrites, thereby promoting nascent dendrite stability. (D) Augmin may be required to maintain the correct polarity of microtubules within dendrites, thereby ensuring the correct motor-driven delivery along microtubules of material required for higher-order dendritic branching.
**Figure 6.** Non-mutually exclusive possibilities for how Augmin may promote **higher-order dendritic branching.** Cartoons representing how Augmin-mediated microtubule nucleation may promote the formation of terminal dendrites from either secondary or tertiary dendrites in the distal region of the class IV da neuron. (A) Augmin-γ-TuRC (purple and pink respectively) may assemble at the base of nascent dendrites and nucleate microtubules at angles that allow the daughter microtubules to grow into and stabilise the nascent dendrite. (B) Augmin-γ-TuRC may be required to amplify microtubules throughout the dendritic arbor, increasing the likelihood that microtubules turn into and stabilise nascent dendrites. (C) Augmin-γ-TuRC may be required to amplify anterograde microtubules within nascent dendrites, thereby promoting nascent dendrite stability. (D) Augmin may be required to maintain the correct polarity of microtubules within dendrites, thereby ensuring the correct motor-driven delivery along microtubules of material required for higher-order dendritic branching.
References

Conduit, P. T., Wainman, A. and Raff, J. W. (2015). Centrosome function and assembly in animal cells. *Nat Rev Mol Cell Bio* **16**, 611–624.

Cunha-Ferreira, I., Chazeau, A., Buijs, R. R., Stucchi, R., Will, L., Pan, X., Adolfs, Y., Meer, C. van der, Wolthuis, J. C., Kahn, O. I., et al. (2018). The HAUS Complex Is a Key Regulator of Non-centrosomal Microtubule Organization during Neuronal Development. *Cell Reports* **24**, 791–800.

Dobbelaere, J., Josué, F., Suijkerbuijk, S., Baum, B., Tapon, N. and Raff, J. (2008). A genome-wide RNAi screen to dissect centriole duplication and centrosome maturation in Drosophila. *Plos Biol* **6**, e224.

Feng, C., Thyagarajan, P., Shorey, M., Seebold, D. Y., Weiner, A. T., Albertson, R. M., Rao, K. S., Sagasti, A., Goetschius, D. J. and Rolls, M. M. (2019). Patronin-mediated minus end growth is required for dendritic microtubule polarity. *J Cell Biology* **218**, 2309–2328.

Fu, M., McAlear, T. S., Nguyen, H., Oses-Prieto, J. A., Valenzuela, A., Shi, R. D., Perrino, J. J., Huang, T.-T., Burlingame, A. L., Bechstedt, S., et al. (2019). The Golgi Outpost Protein TPPP Nucleates Microtubules and Is Critical for Myelination. *Cell* **179**, 132-146.e14.

Gabel, C. A., Li, Z., DeMarco, A. G., Zhang, Z., Yang, J., Hall, M. C., Barford, D. and Chang, L. (2022). Molecular architecture of the augmin complex. *Nat Commun* **13**, 5449.

Gao, F.-B., Brenman, J. E., Jan, L. Y. and Jan, Y. N. (1999). Genes regulating dendritic outgrowth, branching, and routing in Drosophila. *Gene Dev* **13**, 2549–2561.

Garbrecht, J., Laos, T., Holzer, E., Dillinger, M. and Dammermann, A. (2021). An acentriolar centrosome at the C. elegans ciliary base. *Curr Biol* **31**, 2418-2428.e8.

Goshima, G., Wollman, R., Goodwin, S. S., Zhang, N., Scholey, J. M., Vale, R. D. and Stuurman, N. (2007). Genes Required for Mitotic Spindle Assembly in *Drosophila* S2 Cells. *Science* **316**, 417–421.

Goshima, G., Mayer, M., Zhang, N., Stuurman, N. and Vale, R. D. (2008). Augmin: a protein complex required for centrosome-independent microtubule generation within the spindle. *J Cell Biology* **181**, 421–429.

Grueber, W. B., Jan, L. Y. and Jan, Y.-N. (2002). Tiling of the Drosophila epidermis by multidendritic sensory neurons. *Development (Cambridge, England)* **129**, 2867–2878.

Harterink, M., Edwards, S. L., Haan, B. de, Yau, K. W., Heuvel, S. van den, Kapitein, L. C., Miller, K. G. and Hoogenraad, C. C. (2018). Local microtubule organization promotes cargo transport in C. elegans dendrites. *J Cell Sci* **131**, jcs.223107.
References

Conduit, P. T., Wainman, A. and Raff, J. W. (2015). Centrosome function and assembly in animal cells. *Nat Rev Mol Cell Bio* **16**, 611–624.

Cunha-Ferreira, I., Chazeau, A., Buijs, R. R., Stucchi, R., Will, L., Pan, X., Adolfs, Y., Meer, C. van der, Wolthuis, J. C., Kahn, O. I., et al. (2018). The HAUS Complex Is a Key Regulator of Non-centrosomal Microtubule Organization during Neuronal Development. *Cell Reports* **24**, 791–800.

Dobbelaere, J., Josué, F., Suijkerbuijk, S., Baum, B., Tapon, N. and Raff, J. (2008). A genome-wide RNAi screen to dissect centriole duplication and centrosome maturation in Drosophila. *Plos Biol* **6**, e224.

Feng, C., Thyagarajan, P., Shorey, M., Seebold, D. Y., Weiner, A. T., Albertson, R. M., Rao, K. S., Sagasti, A., Goetschius, D. J. and Rolls, M. M. (2019). Patronin-mediated minus end growth is required for dendritic microtubule polarity. *J Cell Biology* **218**, 2309–2328.

Fu, M., McAlear, T. S., Nguyen, H., Oses-Prieto, J. A., Valenzuela, A., Shi, R. D., Perrino, J. J., Huang, T.-T., Burlingame, A. L., Bechstedt, S., et al. (2019). The Golgi Outpost Protein TPPP Nucleates Microtubules and Is Critical for Myelination. *Cell* **179**, 132-146.e14.

Gabel, C. A., Li, Z., DeMarco, A. G., Zhang, Z., Yang, J., Hall, M. C., Barford, D. and Chang, L. (2022). Molecular architecture of the augmin complex. *Nat Commun* **13**, 5449.

Gao, F.-B., Brenman, J. E., Jan, L. Y. and Jan, Y. N. (1999). Genes regulating dendritic outgrowth, branching, and routing in Drosophila. *Gene Dev* **13**, 2549–2561.

Garbrecht, J., Laos, T., Holzer, E., Dillinger, M. and Dammermann, A. (2021). An acentriolar centrosome at the C. elegans ciliary base. *Curr Biol* **31**, 2418-2428.e8.

Goshima, G., Wollman, R., Goodwin, S. S., Zhang, N., Scholey, J. M., Vale, R. D. and Stuurman, N. (2007). Genes Required for Mitotic Spindle Assembly in *Drosophila* S2 Cells. *Science* **316**, 417–421.

Goshima, G., Mayer, M., Zhang, N., Stuurman, N. and Vale, R. D. (2008). Augmin: a protein complex required for centrosome-independent microtubule generation within the spindle. *J Cell Biology* **181**, 421–429.

Grueber, W. B., Jan, L. Y. and Jan, Y.-N. (2002). Tiling of the Drosophila epidermis by multidendritic sensory neurons. *Development (Cambridge, England)* **129**, 2867–2878.

Harterink, M., Edwards, S. L., Haan, B. de, Yau, K. W., Heuvel, S. van den, Kapitein, L. C., Miller, K. G. and Hoogenraad, C. C. (2018). Local microtubule organization promotes cargo transport in C. elegans dendrites. *J Cell Sci* **131**, jcs.223107.
Hu, C., Petersen, M., Hoyer, N., Spitzweck, B., Tenedini, F., Wang, D., Gruschka, A., Burchardt, L. S., Szpotowicz, E., Schweizer, M., et al. (2017). Sensory integration and neuromodulatory feedback facilitate Drosophila mechanosensitive behavior. *Nat. Neurosci.* **20**, 1085–1095.

Jan, Y.-N. and Jan, L. Y. (2010). Branching out: mechanisms of dendritic arborization. *Nat Rev Neurosci* **11**, 316–28.

Janson, M. E., Setty, T. G., Paoletti, A. and Tran, P. T. (2005). Efficient formation of bipolar microtubule bundles requires microtubule-bound γ-tubulin complexes. *J. Cell Biol.* **169**, 297–308.

Kamasaki, T., O'Toole, E., Kita, S., Osumi, M., Usukura, J., McIntosh, J. R. and Goshima, G. (2013). Augmin-dependent microtubule nucleation at microtubule walls in the spindle. *J Cell Biol* **202**, 25–33.

Kapitein, L. C. and Hoogenraad, C. C. (2015). Building the Neuronal Microtubule Cytoskeleton. *Neuron* **87**, 492–506.

Kollman, J. M., Merdes, A., Mourey, L. and Agard, D. A. (2011). Microtubule nucleation by γ-tubulin complexes. *Nat Rev Mol Cell Bio* **12**, 709–721.

Kuo, C. T., Jan, L. Y. and Jan, Y. N. (2005). Dendrite-specific remodeling of Drosophila sensory neurons requires matrix metalloproteases, ubiquitin-proteasome, and ecdysone signaling. *P Natl Acad Sci USA* **102**, 15230–15235.

Lawo, S., Bashkurov, M., Mullin, M., Ferreria, M. G., Kittler, R., Habermann, B., Tagliaferro, A., Poser, I., Hutchins, J. R. A., Hegemann, B., et al. (2009). HAUS, the 8-subunit human Augmin complex, regulates centrosome and spindle integrity. *Curr Biol* **19**, 816–826.

Liu, T., Tian, J., Wang, G., Yu, Y., Wang, C., Ma, Y., Zhang, X., Xia, G., Liu, B. and Kong, Z. (2014). Augmin Triggers Microtubule-Dependent Microtubule Nucleation in Interphase Plant Cells. *Curr Biol* **24**, 2708–2713.

Lüders, J. (2020). Nucleating microtubules in neurons: Challenges and solutions. *Dev Neurobiol*.

Magescas, J., Eskinazi, S., Tran, M. V. and Feldman, J. L. (2021). Centriole-less pericentriolar material serves as a microtubule organizing center at the base of C. elegans sensory cilia. *Curr Biol* **31**, 2410–2417.e6.

Moutinho-Pereira, S., Stuurman, N., Afonso, O., Hornsveld, M., Aguiar, P., Goshima, G., Vale, R. D. and Maiato, H. (2013). Genes involved in centrosome-independent mitotic spindle assembly in Drosophila S2 cells. *Proc. Natl. Acad. Sci.* **110**, 19808–19813.

Mukherjee, A., Brooks, P. S., Bernard, F., Guichet, A. and Conduit, P. T. (2020). Microtubules originate asymmetrically at the somatic Golgi and are guided via Kinesin2 to maintain polarity in neurons. *Elife* **9**, e58943.
Hu, C., Petersen, M., Hoyer, N., Spitzweck, B., Tenedini, F., Wang, D., Gruschka, A., Burchardt, L. S., Szpotowicz, E., Schweizer, M., et al. (2017). Sensory integration and neuromodulatory feedback facilitate Drosophila mechanosensitive behavior. *Nat. Neurosci.* **20**, 1085–1095.

Jan, Y.-N. and Jan, L. Y. (2010). Branching out: mechanisms of dendritic arborization. *Nat Rev Neurosci* **11**, 316–28.

Janson, M. E., Setty, T. G., Paoletti, A. and Tran, P. T. (2005). Efficient formation of bipolar microtubule bundles requires microtubule-bound γ-tubulin complexes. *J. Cell Biol.* **169**, 297–308.

Kamasaki, T., O'Toole, E., Kita, S., Osumi, M., Usukura, J., McIntosh, J. R. and Goshima, G. (2013). Augmin-dependent microtubule nucleation at microtubule walls in the spindle. *J Cell Biol* **202**, 25–33.

Kapitein, L. C. and Hoogenraad, C. C. (2015). Building the Neuronal Microtubule Cytoskeleton. *Neuron* **87**, 492–506.

Kollman, J. M., Merdes, A., Mourey, L. and Agard, D. A. (2011). Microtubule nucleation by γ-tubulin complexes. *Nat Rev Mol Cell Bio* **12**, 709–721.

Kuo, C. T., Jan, L. Y. and Jan, Y. N. (2005). Dendrite-specific remodeling of Drosophila sensory neurons requires matrix metalloproteases, ubiquitin-proteasome, and ecdysone signaling. *P Natl Acad Sci USA* **102**, 15230–15235.

Lawo, S., Bashkurov, M., Mullin, M., Ferreria, M. G., Kittler, R., Habermann, B., Tagliaferro, A., Poser, I., Hutchins, J. R. A., Hegemann, B., et al. (2009). HAUS, the 8-subunit human Augmin complex, regulates centrosome and spindle integrity. *Curr Biol* **19**, 816–826.

Liu, T., Tian, J., Wang, G., Yu, Y., Wang, C., Ma, Y., Zhang, X., Xia, G., Liu, B. and Kong, Z. (2014). Augmin Triggers Microtubule-Dependent Microtubule Nucleation in Interphase Plant Cells. *Curr Biol* **24**, 2708–2713.

Lüders, J. (2020). Nucleating microtubules in neurons: Challenges and solutions. *Dev Neurobiol*.

Magescas, J., Eskinazi, S., Tran, M. V. and Feldman, J. L. (2021). Centriole-less pericentriolar material serves as a microtubule organizing center at the base of C. elegans sensory cilia. *Curr Biol* **31**, 2410–2417.e6.

Moutinho-Pereira, S., Stuurman, N., Afonso, O., Hornsveld, M., Aguiar, P., Goshima, G., Vale, R. D. and Maiato, H. (2013). Genes involved in centrosome-independent mitotic spindle assembly in Drosophila S2 cells. *Proc. Natl. Acad. Sci.* **110**, 19808–19813.

Mukherjee, A., Brooks, P. S., Bernard, F., Guichet, A. and Conduit, P. T. (2020). Microtubules originate asymmetrically at the somatic Golgi and are guided via Kinesin2 to maintain polarity in neurons. *Elife* **9**, e58943.
Murata, T., Sonobe, S., Baskin, T. I., Hyodo, S., Hasezawa, S., Nagata, T., Horio, T. and Hasebe, M. (2005). Microtubule-dependent microtubule nucleation based on recruitment of γ-tubulin in higher plants. *Nat Cell Biol* **7**, 961–968.

Nguyen, M. M., Stone, M. C. and Rolls, M. M. (2011). Microtubules are organized independently of the centrosome in Drosophila neurons. *Neural Dev* **6**, 38.

Nguyen, M. M., McCracken, C. J., Milner, E. S., Goetschius, D. J., Weiner, A. T., Long, M. K., Michael, N. L., Munro, S. and Rolls, M. M. (2014). γ-Tubulin controls neuronal microtubule polarity independently of Golgi outposts. *Mol Biol Cell* **25**, 2039–2050.

Nithianandam, V. and Chien, C.-T. (2018). Actin blobs prefigure dendrite branching sites. *J Cell Biol* **217**, jcb.201711136.

Ori-McKenney, K. M., Jan, L. Y. and Jan, Y.-N. (2012). Golgi outposts shape dendrite morphology by functioning as sites of acentrosomal microtubule nucleation in neurons. *Neuron* **76**, 921–930.

Parrish, J. Z., Xu, P., Kim, C. C., Jan, L. Y. and Jan, Y. N. (2009). The microRNA bantam functions in epithelial cells to regulate scaling growth of dendrite arbors in Drosophila sensory neurons. *Neuron* **63**, 788–802.

Petry, S., Groen, A. C., Ishihara, K., Mitchison, T. J. and Vale, R. D. (2013). Branching microtubule nucleation in Xenopus Egg Extracts Mediated by Augmin and TPX2. *Cell* **152**, 768–777.

Qu, X., Kumar, A., Blockus, H., Waites, C. and Bartolini, F. (2019). Activity-Dependent Nucleation of Dynamic Microtubules at Presynaptic Boutons Controls Neurotransmission. *Curr Biol* **29**, 4231-4240.e5.

Reschen, R. F., Colombié, N., Wheatley, L., Dobbelaere, J., Johnston, D. S., Ohkura, H. and Raff, J. W. (2012). Dgp71WD is required for the assembly of the acentrosomal Meiosis I spindle, and is not a general targeting factor for the γ-TuRC. *Biol Open* **1**, 422–429.

Rolls, M. M. (2011). Neuronal polarity in Drosophila: Sorting out axons and dendrites. *Dev Neurobiol* **71**, 419–429.

Rolls, M. M. and Jegla, T. J. (2015). Neuronal polarity: an evolutionary perspective. *J Exp Biology* **218**, 572–580.

Rolls, M. M., Thyagarajan, P. and Feng, C. (2021). Microtubule dynamics in healthy and injured neurons. *Dev Neurobiol* **81**, 321–332.

Sanchez, A. D. and Feldman, J. L. (2016). Microtubule-organizing centers: from the centrosome to non-centrosomal sites. *Curr Opin Cell Biol* **44**, 93–101.

Sánchez-Huertas, C. and Lüders, J. (2015). The augmin connection in the geometry of microtubule networks. *Curr Biol* **25**, R294–9.
**Murata, T., Sonobe, S., Baskin, T. I., Hyodo, S., Hasezawa, S., Nagata, T., Horio, T. and Hasebe, M.** (2005). Microtubule-dependent microtubule nucleation based on recruitment of γ-tubulin in higher plants. *Nat Cell Biol* **7**, 961–968.

**Nguyen, M. M., Stone, M. C. and Rolls, M. M.** (2011). Microtubules are organized independently of the centrosome in Drosophila neurons. *Neural Dev* **6**, 38.

**Nguyen, M. M., McCracken, C. J., Milner, E. S., Goetschius, D. J., Weiner, A. T., Long, M. K., Michael, N. L., Munro, S. and Rolls, M. M.** (2014). γ-Tubulin controls neuronal microtubule polarity independently of Golgi outposts. *Mol Biol Cell* **25**, 2039–2050.

**Nithianandam, V. and Chien, C.-T.** (2018). Actin blobs prefigure dendrite branching sites. *J Cell Biol* **217**, jcb.201711136.

**Ori-McKenney, K. M., Jan, L. Y. and Jan, Y.-N.** (2012). Golgi outposts shape dendrite morphology by functioning as sites of acentrosomal microtubule nucleation in neurons. *Neuron* **76**, 921–930.

**Parrish, J. Z., Xu, P., Kim, C. C., Jan, L. Y. and Jan, Y. N.** (2009). The microRNA bantam functions in epithelial cells to regulate scaling growth of dendrite arbors in Drosophila sensory neurons. *Neuron* **63**, 788–802.

**Petry, S., Groen, A. C., Ishihara, K., Mitchison, T. J. and Vale, R. D.** (2013). Branching microtubule nucleation in Xenopus Egg Extracts Mediated by Augmin and TPX2. *Cell* **152**, 768–777.

**Qu, X., Kumar, A., Blockus, H., Waites, C. and Bartolini, F.** (2019). Activity-Dependent Nucleation of Dynamic Microtubules at Presynaptic Boutons Controls Neurotransmission. *Curr Biol* **29**, 4231-4240.e5.

**Reschen, R. F., Colombié, N., Wheatley, L., Dobbelaere, J., Johnston, D. S., Ohkura, H. and Raff, J. W.** (2012). Dgp71WD is required for the assembly of the acentrosomal Meiosis I spindle, and is not a general targeting factor for the γ-TuRC. *Biol Open* **1**, 422–429.

**Rolls, M. M.** (2011). Neuronal polarity in Drosophila: Sorting out axons and dendrites. *Dev Neurobiol* **71**, 419–429.

**Rolls, M. M. and Jegla, T. J.** (2015). Neuronal polarity: an evolutionary perspective. *J Exp Biology* **218**, 572–580.

**Rolls, M. M., Thyagarajan, P. and Feng, C.** (2021). Microtubule dynamics in healthy and injured neurons. *Dev Neurobiol* **81**, 321–332.

**Sanchez, A. D. and Feldman, J. L.** (2016). Microtubule-organizing centers: from the centrosome to non-centrosomal sites. *Curr Opin Cell Biol* **44**, 93–101.

**Sánchez-Huertas, C. and Lüders, J.** (2015). The augmin connection in the geometry of microtubule networks. *Curr Biol* **25**, R294–9.
Sánchez-Huertas, C., Freixo, F., Viais, R., Lacasa, C., Soriano, E. and Lüders, J. (2016). Non-centrosomal nucleation mediated by augmin organizes microtubules in post-mitotic neurons and controls axonal microtubule polarity. *Nat Commun* **7**, 12187.

Satoh, D., Suyama, R., Kimura, K. and Uemura, T. (2012). High-resolution in vivo imaging of regenerating dendrites of Drosophila sensory neurons during metamorphosis: local filopodial degeneration and heterotypic dendrite-dendrite contacts. *Genes Cells* **17**, 939–951.

Sears, J. C. and Broihier, H. T. (2016). FoxO regulates microtubule dynamics and polarity to promote dendrite branching in Drosophila sensory neurons. *Dev Biol* **418**, 40–54.

Shimono, K., Fujimoto, A., Tsuyama, T., Yamamoto-Kochi, M., Sato, M., Hattori, Y., Sugimura, K., Usui, T., Kimura, K. and Uemura, T. (2009). Multidendritic sensory neurons in the adult Drosophila abdomen: origins, dendritic morphology, and segment- and age-dependent programmed cell death. *Neural Dev* **4**, 37.

Shree, S., Sutradhar, S., Trottier, O., Tu, Y., Liang, X. and Howard, J. (2022). Dynamic instability of dendrite tips generates the highly branched morphologies of sensory neurons. *Sci. Adv.* **8**, eabn0080.

Song, W., Onishi, M., Jan, L. Y. and Jan, Y. N. (2007). Peripheral multidendritic sensory neurons are necessary for rhythmic locomotion behavior in Drosophila larvae. *Proc. Natl. Acad. Sci.* **104**, 5199–5204.

Song, J.-G., King, M. R., Zhang, R., Kadzik, R. S., Thawani, A. and Petry, S. (2018). Mechanism of how augmin directly targets the γ-tubulin ring complex to microtubules. *J Cell Biol* **217**, jcb.201711090.

Stiess, M., Maghelli, N., Kapitein, L. C., Gomis-Rüth, S., Wilsch-Bräuninger, M., Hoogenraad, C. C., Tolić-Nørrelykke, I. M. and Bradke, F. (2010). Axon Extension Occurs Independently of Centrosomal Microtubule Nucleation. *Science* **327**, 704–707.

Stürner, T., Tatarnikova, A., Mueller, J., Schaffran, B., Cuntz, H., Zhang, Y., Nemethova, M., Bogdan, S., Small, V. and Tavosanis, G. (2019). Transient localization of the Arp2/3 complex initiates neuronal dendrite branching in vivo. *Development* **146**, dev171397.

Tas, R. P., Chazeau, A., Cloin, B. M. C., Lambers, M. L. A., Hoogenraad, C. C. and Kapitein, L. C. (2017). Differentiation between Oppositely Oriented Microtubules Controls Polarized Neuronal Transport. *Neuron* **96**, 1264–1271.e5.

Thyagarajan, P., Feng, C., Lee, D., Shorey, M. and Rolls, M. M. (2022). Microtubule polarity is instructive for many aspects of neuronal polarity. *Dev Biol* **486**, 56–70.

Tillery, M., Blake-Hedges, C., Zheng, Y., Buchwalter, R. and Megraw, T. (2018). Centrosomal and Non-Centrosomal Microtubule-Organizing Centers (MTOCs) in Drosophila melanogaster. *Cells* **7**, 121.
Sánchez-Huertas, C., Freixo, F., Viais, R., Lacasa, C., Soriano, E. and Lüders, J. (2016). Non-centrosomal nucleation mediated by augmin organizes microtubules in post-mitotic neurons and controls axonal microtubule polarity. *Nat Commun* **7**, 12187.

Satoh, D., Suyama, R., Kimura, K. and Uemura, T. (2012). High-resolution in vivo imaging of regenerating dendrites of Drosophila sensory neurons during metamorphosis: local filopodial degeneration and heterotypic dendrite-dendrite contacts. *Genes Cells* **17**, 939–951.

Sears, J. C. and Broihier, H. T. (2016). FoxO regulates microtubule dynamics and polarity to promote dendrite branching in Drosophila sensory neurons. *Dev Biol* **418**, 40–54.

Shimono, K., Fujimoto, A., Tsuyama, T., Yamamoto-Kochi, M., Sato, M., Hattori, Y., Sugimura, K., Usui, T., Kimura, K. and Uemura, T. (2009). Multidendritic sensory neurons in the adult Drosophila abdomen: origins, dendritic morphology, and segment- and age-dependent programmed cell death. *Neural Dev* **4**, 37.

Shree, S., Sutradhar, S., Trottier, O., Tu, Y., Liang, X. and Howard, J. (2022). Dynamic instability of dendrite tips generates the highly branched morphologies of sensory neurons. *Sci. Adv.* **8**, eabn0080.

Song, W., Onishi, M., Jan, L. Y. and Jan, Y. N. (2007). Peripheral multidendritic sensory neurons are necessary for rhythmic locomotion behavior in Drosophila larvae. *Proc. Natl. Acad. Sci.* **104**, 5199–5204.

Song, J.-G., King, M. R., Zhang, R., Kadzik, R. S., Thawani, A. and Petry, S. (2018). Mechanism of how augmin directly targets the γ-tubulin ring complex to microtubules. *J Cell Biol* **217**, jcb.201711090.

Stiess, M., Maghelli, N., Kapitein, L. C., Gomis-Rüth, S., Wilsch-Bräuninger, M., Hoogenraad, C. C., Tolić-Nørrelykke, I. M. and Bradke, F. (2010). Axon Extension Occurs Independently of Centrosomal Microtubule Nucleation. *Science* **327**, 704–707.

Stürner, T., Tatarnikova, A., Mueller, J., Schaffran, B., Cuntz, H., Zhang, Y., Nemethova, M., Bogdan, S., Small, V. and Tavosanis, G. (2019). Transient localization of the Arp2/3 complex initiates neuronal dendrite branching in vivo. *Development* **146**, dev171397.

Tas, R. P., Chazeau, A., Cloin, B. M. C., Lambers, M. L. A., Hoogenraad, C. C. and Kapitein, L. C. (2017). Differentiation between Oppositely Oriented Microtubules Controls Polarized Neuronal Transport. *Neuron* **96**, 1264–1271.e5.

Thyagarajan, P., Feng, C., Lee, D., Shorey, M. and Rolls, M. M. (2022). Microtubule polarity is instructive for many aspects of neuronal polarity. *Dev Biol* **486**, 56–70.

Tillery, M., Blake-Hedges, C., Zheng, Y., Buchwalter, R. and Megraw, T. (2018). Centrosomal and Non-Centrosomal Microtubule-Organizing Centers (MTOCs) in Drosophila melanogaster. *Cells* **7**, 121.
Tovey, C. A. and Conduit, P. T. (2018). Microtubule nucleation by γ-tubulin complexes and beyond. *Essays Biochem* **91**, EBC20180028.

Tovey, C. A., Tubman, C. E., Hamrud, E., Zhu, Z., Dyas, A. E., Butterfield, A. N., Fyfe, A., Johnson, E. and Conduit, P. T. (2018). γ-TuRC Heterogeneity Revealed by Analysis of Mozart1. *Curr Biol* **28**, 2314–2323.e6.

Vaadia, R. D., Li, W., Voleti, V., Singhania, A., Hillman, E. M. C. and Grueber, W. B. (2019). Characterization of Proprioceptive System Dynamics in Behaving Drosophila Larvae Using High-Speed Volumetric Microscopy. *Curr. Biol.* **29**, 935-944.e4.

Valenzuela, A., Meservey, L., Nguyen, H. and Fu, M. (2020). Golgi Outposts Nucleate Microtubules in Cells with Specialized Shapes. *Trends Cell Biol*.

Verma, V. and Maresca, T. J. (2019). Direct observation of branching MT nucleation in living animal cells. *J Cell Biology* **218**, 2829–2840.

Vérollet, C., Colombié, N., Daubon, T., Bourbon, H.-M., Wright, M. and Raynaud-Messina, B. (2006). Drosophila melanogaster γ-TuRC is dispensable for targeting γ-tubulin to the centrosome and microtubule nucleation. *J Cell Biology* **172**, 517–528.

Wasteneys, G. O. and Williamson, R. E. (1989). Reassembly of microtubules in Nitella tasmanica: assembly of cortical microtubules in branching clusters and its relevance to steady-state microtubule assembly. *J. Cell Sci.* **93**, 705–714.

Weiner, A. T., Seebold, D. Y., Torres-Gutierrez, P., Folker, C., Swope, R. D., Kothe, G. O., Stoltz, J. G., Zalenski, M. K., Kozlowski, C., Barbera, D. J., et al. (2020). Endosomal Wnt signaling proteins control microtubule nucleation in dendrites. *Plos Biol* **18**, e3000647.

Williams, D. W. and Truman, J. W. (2005). Cellular mechanisms of dendrite pruning in Drosophila: insights from in vivo time-lapse of remodeling dendritic arborizing sensory neurons. *Development* **132**, 3631–3642.

Xiang, Y., Yuan, Q., Vogt, N., Looger, L. L., Jan, L. Y. and Jan, Y. N. (2010). Light-avoidance-mediating photoreceptors tile the Drosophila larval body wall. *Nature* **468**, 921–926.

Xu, Y., Wang, B., Bush, I., Saunders, H. A., Wildonger, J. and Han, C. (2023). Light-induced trapping of endogenous proteins reveals spatiotemporal roles of microtubule and kinesin-1 in dendrite patterning of Drosophila sensory neurons. *bioRxiv* 2023.09.30.560303.

Yalgin, C., Ebrahimi, S., Delandre, C., Yoong, L. F., Akimoto, S., Tran, H., Amikura, R., Spokony, R., Torben-Nielsen, B., White, K. P., et al. (2015). Centrosomin represses dendrite branching by orienting microtubule nucleation. *Nat Neurosci* **18**, 1437–1445.

Yang, S. Z. and Wildonger, J. (2020). Golgi Outposts Locally Regulate Microtubule Orientation in Neurons but Are Not Required for the Overall Polarity of the Dendritic Cytoskeleton. *Genetics* **215**, 435–447.
Tovey, C. A. and Conduit, P. T. (2018). Microtubule nucleation by γ-tubulin complexes and beyond. *Essays Biochem* **91**, EBC20180028.

Tovey, C. A., Tubman, C. E., Hamrud, E., Zhu, Z., Dyas, A. E., Butterfield, A. N., Fyfe, A., Johnson, E. and Conduit, P. T. (2018). γ-TuRC Heterogeneity Revealed by Analysis of Mozart1. *Curr Biol* **28**, 2314–2323.e6.

Vaadia, R. D., Li, W., Voleti, V., Singhania, A., Hillman, E. M. C. and Grueber, W. B. (2019). Characterization of Proprioceptive System Dynamics in Behaving Drosophila Larvae Using High-Speed Volumetric Microscopy. *Curr. Biol.* **29**, 935-944.e4.

Valenzuela, A., Meservey, L., Nguyen, H. and Fu, M. (2020). Golgi Outposts Nucleate Microtubules in Cells with Specialized Shapes. *Trends Cell Biol*.

Verma, V. and Maresca, T. J. (2019). Direct observation of branching MT nucleation in living animal cells. *J Cell Biology* **218**, 2829–2840.

Vérollet, C., Colombié, N., Daubon, T., Bourbon, H.-M., Wright, M. and Raynaud-Messina, B. (2006). Drosophila melanogaster γ-TuRC is dispensable for targeting γ-tubulin to the centrosome and microtubule nucleation. *J Cell Biology* **172**, 517–528.

Wasteneys, G. O. and Williamson, R. E. (1989). Reassembly of microtubules in Nitella tasmanica: assembly of cortical microtubules in branching clusters and its relevance to steady-state microtubule assembly. *J. Cell Sci.* **93**, 705–714.

Weiner, A. T., Seebold, D. Y., Torres-Gutierrez, P., Folker, C., Swope, R. D., Kothe, G. O., Stoltz, J. G., Zalenski, M. K., Kozlowski, C., Barbera, D. J., et al. (2020). Endosomal Wnt signaling proteins control microtubule nucleation in dendrites. *Plos Biol* **18**, e3000647.

Williams, D. W. and Truman, J. W. (2005). Cellular mechanisms of dendrite pruning in Drosophila: insights from in vivo time-lapse of remodeling dendritic arborizing sensory neurons. *Development* **132**, 3631–3642.

Xiang, Y., Yuan, Q., Vogt, N., Looger, L. L., Jan, L. Y. and Jan, Y. N. (2010). Light-avoidance-mediating photoreceptors tile the Drosophila larval body wall. *Nature* **468**, 921–926.

Xu, Y., Wang, B., Bush, I., Saunders, H. A., Wildonger, J. and Han, C. (2023). Light-induced trapping of endogenous proteins reveals spatiotemporal roles of microtubule and kinesin-1 in dendrite patterning of Drosophila sensory neurons. *bioRxiv* 2023.09.30.560303.

Yalgin, C., Ebrahimi, S., Delandre, C., Yoong, L. F., Akimoto, S., Tran, H., Amikura, R., Spokony, R., Torben-Nielsen, B., White, K. P., et al. (2015). Centrosomin represses dendrite branching by orienting microtubule nucleation. *Nat Neurosci* **18**, 1437–1445.

Yang, S. Z. and Wildonger, J. (2020). Golgi Outposts Locally Regulate Microtubule Orientation in Neurons but Are Not Required for the Overall Polarity of the Dendritic Cytoskeleton. *Genetics* **215**, 435–447.
Yoong, L.-F., Lim, H.-K., Tran, H., Lackner, S., Zheng, Z., Hong, P. and Moore, A. W. (2020).
Atypical Myosin Tunes Dendrite Arbor Subdivision. *Neuron*.

Zhong, L., Hwang, R. Y. and Tracey, W. D. (2010). Pickpocket Is a DEG/ENaC Protein Required for Mechanical Nociception in Drosophila Larvae. *Curr. Biol.* **20**, 429–434.

Zhou, W., Chang, J., Wang, X., Savelieff, M. G., Zhao, Y., Ke, S. and Ye, B. (2014). GM130 is required for compartmental organization of dendritic golgi outposts. *Curr Biol* **24**, 1227–1233.

Zhu, Z., Becam, I., Tovey, C. A., Elfarkouchi, A., Yen, E. C., Bernard, F., Guichet, A. and **Conduit, P. T.** (2023). Multifaceted modes of γ-tubulin complex recruitment and microtubule nucleation at mitotic centrosomes. *J. Cell Biol.* **222**, e202212043.

Zupa, E., Würtz, M., Neuner, A., Hoffmann, T., Rettel, M., Böhler, A., Vermeulen, B. J. A., Eustermann, S., Schiebel, E. and Pfeffer, S. (2022). The augmin complex architecture reveals structural insights into microtubule branching. *Nat. Commun.* **13**, 5635.
Yoong, L.-F., Lim, H.-K., Tran, H., Lackner, S., Zheng, Z., Hong, P. and Moore, A. W. (2020). Atypical Myosin Tunes Dendrite Arbor Subdivision. *Neuron*.

Zhong, L., Hwang, R. Y. and Tracey, W. D. (2010). Pickpocket Is a DEG/ENaC Protein Required for Mechanical Nociception in Drosophila Larvae. *Curr. Biol.* **20**, 429–434.

Zhou, W., Chang, J., Wang, X., Savelieff, M. G., Zhao, Y., Ke, S. and Ye, B. (2014). GM130 is required for compartmental organization of dendritic golgi outposts. *Curr Biol* **24**, 1227–1233.

Zhu, Z., Becam, I., Tovey, C. A., Elfarkouchi, A., Yen, E. C., Bernard, F., Guichet, A. and **Conduit, P. T.** (2023). Multifaceted modes of γ-tubulin complex recruitment and microtubule nucleation at mitotic centrosomes. *J. Cell Biol.* **222**, e202212043.

Zupa, E., Würtz, M., Neuner, A., Hoffmann, T., Rettel, M., Böhler, A., Vermeulen, B. J. A., Eustermann, S., Schiebel, E. and Pfeffer, S. (2022). The augmin complex architecture reveals structural insights into microtubule branching. *Nat. Commun.* **13**, 5635.
<figure><img src="image_4.png" /><figcaption>Figure 1</figcaption></figure>

<figure><img src="image_5.png" /></figure>

<figure><img src="image_6.png" /></figure>

<figure><img src="image_7.png" /></figure>

<figure><img src="image_8.png" /></figure>

<figure><img src="image_9.png" /></figure>

<figure><img src="image_10.png" /></figure>

<figure><img src="image_11.png" /></figure>

<figure><img src="image_12.png" /></figure>

<figure><img src="image_13.png" /></figure>

<figure><img src="image_14.png" /></figure>
<figure><img src="image_4.png" /><figcaption>Figure 1</figcaption></figure>

developing class IV v'ada neurons

<figure><img src="image_5.png" /></figure>

<figure><img src="image_6.png" /></figure>

<figure><img src="image_7.png" /></figure>

<figure><img src="image_8.png" /></figure>

<figure><img src="image_9.png" /></figure>

<figure><img src="image_10.png" /></figure>

<figure><img src="image_11.png" /></figure>

<figure><img src="image_12.png" /></figure>

<figure><img src="image_13.png" /></figure>

<figure><img src="image_14.png" /></figure>
<figure><img src="image_15.png" /><figcaption>Figure 2</figcaption></figure>
<figure><img src="image_15.png" /><figcaption>Figure 2</figcaption></figure>
Figure 3

A

control-RNAi

<figure><img src="image_16.png" /></figure>

grip71<sup>NEDD1</sup>-RNAi2

<figure><img src="image_17.png" /></figure>

dgt2-RNAi2

<figure><img src="image_18.png" /></figure>

dgt3-RNAi1

<figure><img src="image_19.png" /></figure>

dgt4-RNAi

<figure><img src="image_20.png" /></figure>

dgt5-RNAi

<figure><img src="image_21.png" /></figure>

B

perimeter area 72h APF

<figure><img src="image_22.png" /></figure>

C

tip number 72h APF

<figure><img src="image_23.png" /></figure>

D

primary
dendrite length

<figure><img src="image_24.png" /></figure>

E

primary/secondary
dendrite length

<figure><img src="image_25.png" /></figure>

F

primary/secondary
branch points

<figure><img src="image_26.png" /></figure>

G

length / major
branchpoint ratio

<figure><img src="image_27.png" /></figure>
Figure 3

A

control-RNAi

<figure><img src="image_16.png" /></figure>

grip71<sup>NEDD1</sup>-RNAi2

<figure><img src="image_17.png" /></figure>

dgt2-RNAi2

<figure><img src="image_18.png" /></figure>

dgt3-RNAi1

<figure><img src="image_19.png" /></figure>

dgt4-RNAi

<figure><img src="image_20.png" /></figure>

dgt5-RNAi

<figure><img src="image_21.png" /></figure>

B

perimeter area 72h APF

<figure><img src="image_22.png" /></figure>

C

tip number 72h APF

<figure><img src="image_23.png" /></figure>

D

primary
dendrite length

<figure><img src="image_24.png" /></figure>

E

primary/secondary
dendrite length

<figure><img src="image_25.png" /></figure>

F

primary/secondary
branch points

<figure><img src="image_26.png" /></figure>

G

length / major
branchpoint ratio

<figure><img src="image_27.png" /></figure>
<figure><img src="image_28.png" /><figcaption>Figure 4</figcaption></figure>
<figure><img src="image_28.png" /><figcaption>Figure 4</figcaption></figure>
<figure><img src="image_29.png" /><figcaption>Figure 5</figcaption></figure>
<figure><img src="image_29.png" /><figcaption>Figure 5</figcaption></figure>
Figure 6

A

B

C

D

<figure><img src="image_30.png" /></figure>

<figure><img src="image_31.png" /></figure>
Figure 6

A

B

C

D

<figure><img src="image_30.png" /></figure>

<figure><img src="image_31.png" /></figure>
Figure S1

<figure><img src="image_32.png" /></figure>

**Figure S1.** Images to show which dendrites were identified as primary, secondary, tertiary and terminal dendrites. (A) Confocal images show the same pupal class IV v'ada neuron expressing CD4-tdGFP at 72h APF. The image on the right is overlaid with lines indicating primary (blue) and secondary (magenta) dendrites. The distinction between primary and secondary dendrites was made based on the thickness and length of the dendrites and on the angle at which the dendrites emerge from the branchpoint. (B) Confocal images show the same larval class IV ddaC neuron expressing CD4-tdGFP at 76h AEL. The image on the right is overlaid with lines indicating primary (blue), secondary (magenta), tertiary (orange), and terminal (green) dendrites. The distinction was made based on the thickness and length of the dendrites and the angle at which the dendrites emerge from the branchpoint.
Figure S1

<figure><img src="image_32.png" /></figure>

**Figure S1.** Images to show which dendrites were identified as primary, secondary, tertiary and terminal dendrites. (A) Confocal images show the same pupal class IV v'ada neuron expressing CD4-tdGFP at 72h APF. The image on the right is overlaid with lines indicating primary (blue) and secondary (magenta) dendrites. The distinction between primary and secondary dendrites was made based on the thickness and length of the dendrites and on the angle at which the dendrites emerge from the branchpoint. (B) Confocal images show the same larval class IV ddaC neuron expressing CD4-tdGFP at 76h AEL. The image on the right is overlaid with lines indicating primary (blue), secondary (magenta), tertiary (orange), and terminal (green) dendrites. The distinction was made based on the thickness and length of the dendrites and the angle at which the dendrites emerge from the branchpoint.
Figure S2

<figure><img src="image_33.png" /></figure>

**Figure S2. RT-qPCR analysis of wing disks expressing either γ-TuRC-RNAi or Augmin-RNAi.** Graph shows mRNA levels relative to control estimated using quantitative RT-PCR for grip91 and dgt4 in imaginal discs expressing grip91-RNAi and dgt4-RNAi, respectively. For each condition, 3 biological replicate samples (each 50 discs from 25 flies) of mRNA were prepared. For each biological replicate, 3 repeats of cDNA preparation and subsequent qPCR analysis were performed to generate the 9 values that appear on the graph. Each repeat was normalised to a control value of 1 obtained from discs of w1118 flies by using values from 4 housekeeping genes. The list of genes and the sequences of the primers used are shown in Table S3. Raw data can be found in Source data file 1. The results are presented as the mean 95% CIs. One-sample t-tests were used to test for a significance difference from 1. P values are shown above each dataset.
Figure S2

<figure><img src="image_33.png" /></figure>

**Figure S2. RT-qPCR analysis of wing disks expressing either γ-TuRC-RNAi or Augmin-RNAi.** Graph shows mRNA levels relative to control estimated using quantitative RT-PCR for grip91 and dgt4 in imaginal discs expressing grip91-RNAi and dgt4-RNAi, respectively. For each condition, 3 biological replicate samples (each 50 discs from 25 flies) of mRNA were prepared. For each biological replicate, 3 repeats of cDNA preparation and subsequent qPCR analysis were performed to generate the 9 values that appear on the graph. Each repeat was normalised to a control value of 1 obtained from discs of w1118 flies by using values from 4 housekeeping genes. The list of genes and the sequences of the primers used are shown in Table S3. Raw data can be found in Source data file 1. The results are presented as the mean 95% CIs. One-sample t-tests were used to test for a significance difference from 1. P values are shown above each dataset.
Figure S3

<figure><img src="image_34.png" /><figcaption>neurons expressing γ-tubulin-GFP + grip91-RNAi1 in class I (ddaE/ddaD) neurons</figcaption></figure>

<figure><img src="image_35.png" /></figure>

G Quantifying γ-tub23C-sfGFP signal at the somatic Golgi in class I da neurons

<figure><img src="image_36.png" /></figure>

H control neurons

<figure><img src="image_37.png" /></figure>

I grip91<sup>GCP3</sup>-RNAi1 neurons

<figure><img src="image_38.png" /></figure>
Figure S3

<figure><img src="image_34.png" /><figcaption>neurons expressing γ-tubulin-GFP + grip91-RNAi1 in class I (ddaE/ddaD) neurons</figcaption></figure>

<figure><img src="image_35.png" /></figure>

G Quantifying γ-tub23C-sfGFP signal at the somatic Golgi in class I da neurons

<figure><img src="image_36.png" /></figure>

H control neurons

<figure><img src="image_37.png" /></figure>

I grip91<sup>GCP3</sup>-RNAi1 neurons

<figure><img src="image_38.png" /></figure>
**Figure S3. Depletion of Grip91<sup>GCP3</sup> results in loss of γ-tubulin-GFP from somatic Golgi stacks. (A-F)** Confocal images show the somas of sensory neurons within the dorsal cluster of 3rd instar larva expressing endogenously-tagged γ-tubulin-GFP alone (A-C) or γ-tubulin-GFP and UAS-grip91<sup>GCP3</sup>-RNAi (expressed by 221-Gal4 specifically within class I ddaD and ddaE neurons) (D-F) and immunostained for GFP (green), HRP (marking membranes including Golgi stacks, greyscale), and GM130 (marking cis-Golgi, magenta). (A, A') and (D, D') show a wide view of the dorsal clusters, with (A) and (D) showing the overlay of the γ-tubulin-GFP and HRP signals and (A') and (D') showing just the γ-tubulin-GFP signal. (B), (C), (E), and (F) display class I ddaE neurons (B,E), which express the RNAi constructs, and class IV ddaC neurons (C,F), which do not express the RNAi constructs. Note how γ-tubulin-GFP associates with HRP and GM130 in da neurons under normal conditions (A-C) but does not associate with HRP and GM130 in class I da neurons when grip91<sup>GCP3</sup>-RNAi is expressed (D,E). Note also how γ-tubulin-GFP still associates with Golgi within both other da and ES neurons that do not express grip91<sup>GCP3</sup>-RNAi (D,F). Scale bars are indicated within the images. (G) Graph shows fluorescence intensity measurements of somatic Golgi stacks within either class I da neurons expressing γ-tubulin-GFP alone (grey, n=218 Golgi stacks from 17 neurons) or class I da neurons expressing both γ-tubulin-GFP and grip91<sup>GCP3</sup>-RNAi (red, n=249 Golgi stacks from 17 neurons). P<0.0001, unpaired two-sided t-test on log₁₀ values. (H, I) Images showing examples of class I ddaE neuron soma from 3rd instar larva expressing endogenously-tagged γ-tubulin-GFP alone (H, control neurons) or γ-tubulin-GFP and UAS-grip91<sup>GCP3</sup>-RNAi (I, grip91<sup>GCP3</sup>-RNAi neurons) and immunostained for GFP (green) and HRP (greyscale). Note how the γ-tubulin-GFP signal is predominantly concentrated at Golgi stacks in control neurons but is dispersed throughout the cytosol in grip91<sup>GCP3</sup>-RNAi neurons.
**Figure S3. Depletion of Grip91<sup>GCP3</sup> results in loss of γ-tubulin-GFP from somatic Golgi stacks. (A-F)** Confocal images show the somas of sensory neurons within the dorsal cluster of 3rd instar larva expressing endogenously-tagged γ-tubulin-GFP alone (A-C) or γ-tubulin-GFP and UAS-grip91<sup>GCP3</sup>-RNAi (expressed by 221-Gal4 specifically within class I ddaD and ddaE neurons) (D-F) and immunostained for GFP (green), HRP (marking membranes including Golgi stacks, greyscale), and GM130 (marking cis-Golgi, magenta). (A, A') and (D, D') show a wide view of the dorsal clusters, with (A) and (D) showing the overlay of the γ-tubulin-GFP and HRP signals and (A') and (D') showing just the γ-tubulin-GFP signal. (B), (C), (E), and (F) display class I ddaE neurons (B,E), which express the RNAi constructs, and class IV ddaC neurons (C,F), which do not express the RNAi constructs. Note how γ-tubulin-GFP associates with HRP and GM130 in da neurons under normal conditions (A-C) but does not associate with HRP and GM130 in class I da neurons when grip91<sup>GCP3</sup>-RNAi is expressed (D,E). Note also how γ-tubulin-GFP still associates with Golgi within both other da and ES neurons that do not express grip91<sup>GCP3</sup>-RNAi (D,F). Scale bars are indicated within the images. (G) Graph shows fluorescence intensity measurements of somatic Golgi stacks within either class I da neurons expressing γ-tubulin-GFP alone (grey, n=218 Golgi stacks from 17 neurons) or class I da neurons expressing both γ-tubulin-GFP and grip91<sup>GCP3</sup>-RNAi (red, n=249 Golgi stacks from 17 neurons). P<0.0001, unpaired two-sided t-test on log₁₀ values. (H, I) Images showing examples of class I ddaE neuron soma from 3rd instar larva expressing endogenously-tagged γ-tubulin-GFP alone (H, control neurons) or γ-tubulin-GFP and UAS-grip91<sup>GCP3</sup>-RNAi (I, grip91<sup>GCP3</sup>-RNAi neurons) and immunostained for GFP (green) and HRP (greyscale). Note how the γ-tubulin-GFP signal is predominantly concentrated at Golgi stacks in control neurons but is dispersed throughout the cytosol in grip91<sup>GCP3</sup>-RNAi neurons.
<figure><img src="image_39.png" /></figure>

A example of class IV > class I

<figure><img src="image_40.png" /></figure>

B example of class IV = class I

<figure><img src="image_41.png" /></figure>

<figure><img src="image_42.png" /></figure>

<figure><img src="image_43.png" /></figure>

C dgt5-Gal4 driving UAS-myr-GFP
n = 35 clusters analysed

<figure><img src="image_44.png" /></figure>

Figure S4. The Augmin component Dgt5 is expressed at higher levels in class IV neurons compared to class I neurons. (A,B) Confocal images show the somas of sensory neurons within the dorsal cluster of 3rd instar larva expressing UAS-Myr-GFP (green) under the control of a dgt5-Gal4trap and immunostained for HRP (marking neuronal membranes, magenta). Yellow dotted lines on the dgt5-Gal4trap images show the outlines of the neuronal somas identified with HRP staining. Based on neuronal morphology and position within the cluster, class I ddaE and ddaD, class IV ddaC, and external sensory neurons could be identified and are indicated. The panels on the left and right show examples of where dgt5-Gal4trap expression is higher in the class IV ddaC neuron compared to the class I neurons or roughly equal, respectively. (C) Graph quantifying the frequency of observed differences or similarities in expression levels of dgt5-Gal4trap between class I and class IV neurons from 35 clusters. Note that expression is normally higher in class IV neurons than class I neurons.
Figure S4

A example of class IV > class I

<figure><img src="image_39.png" /></figure>

B example of class IV = class I

<figure><img src="image_40.png" /></figure>

<figure><img src="image_41.png" /></figure>

<figure><img src="image_42.png" /></figure>

C dgt5-Gal4 driving UAS-myr-GFP
n = 35 clusters analysed

<figure><img src="image_43.png" /></figure>

Figure S4. The Augmin component Dgt5 is expressed at higher levels in class IV neurons compared to class I neurons. (A,B) Confocal images show the somas of sensory neurons within the dorsal cluster of 3rd instar larva expressing UAS-Myr-GFP (green) under the control of a dgt5-Gal4trap and immunostained for HRP (marking neuronal membranes, magenta). Yellow dotted lines on the dgt5-Gal4trap images show the outlines of the neuronal somas identified with HRP staining. Based on neuronal morphology and position within the cluster, class I ddaE and ddaD, class IV ddaC, and external sensory neurons could be identified and are indicated. The panels on the left and right show examples of where dgt5-Gal4trap expression is higher in the class IV ddaC neuron compared to the class I neurons or roughly equal, respectively. (C) Graph quantifying the frequency of observed differences or similarities in expression levels of dgt5-Gal4trap between class I and class IV neurons from 35 clusters. Note that expression is normally higher in class IV neurons than class I neurons.
<table><caption>Table S1</caption><thead><tr><th>lab code</th><th>gene</th><th>RNAi type</th><th>stock no.</th><th>40D present</th></tr></thead><tbody><tr><td>R45</td><td>g-tub37C</td><td>TRiP valium 20</td><td>BL32513</td><td>n/a</td></tr><tr><td>R1</td><td>g-tub23C RNAi1</td><td>TRiP valium 1</td><td>BL31204</td><td>n/a</td></tr><tr><td>R21</td><td>g-tub23C RNAi2</td><td>VDRC GD</td><td>v19130</td><td>n/a</td></tr><tr><td>R33</td><td>grip84<sup>GCP2</sup> RNAi1</td><td>VDRC KK</td><td>v105640</td><td>no</td></tr><tr><td>R41</td><td>grip84<sup>GCP2</sup> RNAi2</td><td>TRiP valium 20</td><td>BL33548</td><td>n/a</td></tr><tr><td>R34</td><td>grip91<sup>GCP3</sup> RNAi1</td><td>VDRC KK</td><td>v104667</td><td>no</td></tr><tr><td>R42</td><td>grip91<sup>GCP3</sup> RNAi2</td><td>TRiP valium 1</td><td>BL31201</td><td>n/a</td></tr></tbody></table>

<table><caption>Table S2</caption><thead><tr><th>lab code</th><th>gene</th><th>RNAi type</th><th>stock no.</th><th>40D present</th></tr></thead><tbody><tr><td>R65</td><td>grip71<sup>NEDD1</sup> RNAi1</td><td>VDRC GD</td><td>v31228</td><td>n/a</td></tr><tr><td>R74</td><td>grip71<sup>NEDD1</sup> RNAi2</td><td>NIG</td><td>10346R-1</td><td>n/a</td></tr><tr><td>R64</td><td>dgt2 RNAi1</td><td>TRiP valium 1</td><td>BL31729</td><td>n/a</td></tr><tr><td>R55</td><td>dgt2 RNAi2</td><td>VDRC TRiP valium 20</td><td>v330504</td><td>n/a</td></tr><tr><td>R68</td><td>dgt3 RNAi1</td><td>VDRC KK</td><td>103980</td><td>no</td></tr><tr><td>R62</td><td>dgt3 RNAi2</td><td>VDRC TRiP valium 20</td><td>BL58137</td><td>n/a</td></tr><tr><td>R68</td><td>dgt4 RNAi</td><td>VDRC KK</td><td>v108969</td><td>no</td></tr><tr><td>R63</td><td>dgt5 RNAi</td><td>VDRC TRiP valium 20</td><td>BL60366</td><td>n/a</td></tr></tbody></table>

<table><caption>Table S3</caption><thead><tr><th>Gene</th><th>Forward primer (5'-&gt;3')</th><th>Reverse primer (5'-&gt;3')</th></tr></thead><tbody><tr><td>a-tub84b</td><td>CACACCACCCTGGAGCATTC</td><td>CCAATCAGACGGTTCAGGTTG</td></tr><tr><td>SOP</td><td>CACCCCAATAAAGTTGATAGACCT</td><td>ACCACCACGAGAGCCAAAT</td></tr><tr><td>RPL32</td><td>CTTCATCCGCCACCAGTC</td><td>CGACGCACTCTGTTGTCG</td></tr><tr><td>GAPDH</td><td>GATGCTCCCATGTTCGTTTGC</td><td>GTCCATCACGCCACAACTTG</td></tr><tr><td>dgt4</td><td>CGCCATCATTGACAACTGAGG</td><td>GCTGTCTTCCTGAAAACGGC</td></tr><tr><td>grip91</td><td>TGTGGCAACCAATTCCACTGA</td><td>TTTTCGTGAACTTCGGGCATC</td></tr></tbody></table>
<table><caption>Table S1</caption><thead><tr><th>lab code</th><th>gene</th><th>RNAi type</th><th>stock no.</th><th>40D present</th></tr></thead><tbody><tr><td>R45</td><td>g-tub37C</td><td>TRiP valium 20</td><td>BL32513</td><td>n/a</td></tr><tr><td>R1</td><td>g-tub23C RNAi1</td><td>TRiP valium 1</td><td>BL31204</td><td>n/a</td></tr><tr><td>R21</td><td>g-tub23C RNAi2</td><td>VDRC GD</td><td>v19130</td><td>n/a</td></tr><tr><td>R33</td><td>grip84<sup>GCP2</sup> RNAi1</td><td>VDRC KK</td><td>v105640</td><td>no</td></tr><tr><td>R41</td><td>grip84<sup>GCP2</sup> RNAi2</td><td>TRiP valium 20</td><td>BL33548</td><td>n/a</td></tr><tr><td>R34</td><td>grip91<sup>GCP3</sup> RNAi1</td><td>VDRC KK</td><td>v104667</td><td>no</td></tr><tr><td>R42</td><td>grip91<sup>GCP3</sup> RNAi2</td><td>TRiP valium 1</td><td>BL31201</td><td>n/a</td></tr></tbody></table>

<table><caption>Table S2</caption><thead><tr><th>lab code</th><th>gene</th><th>RNAi type</th><th>stock no.</th><th>40D present</th></tr></thead><tbody><tr><td>R65</td><td>grip71<sup>NEDD1</sup> RNAi1</td><td>VDRC GD</td><td>v31228</td><td>n/a</td></tr><tr><td>R74</td><td>grip71<sup>NEDD1</sup> RNAi2</td><td>NIG</td><td>10346R-1</td><td>n/a</td></tr><tr><td>R64</td><td>dgt2 RNAi1</td><td>TRiP valium 1</td><td>BL31729</td><td>n/a</td></tr><tr><td>R55</td><td>dgt2 RNAi2</td><td>VDRC TRiP valium 20</td><td>v330504</td><td>n/a</td></tr><tr><td>R68</td><td>dgt3 RNAi1</td><td>VDRC KK</td><td>103980</td><td>no</td></tr><tr><td>R62</td><td>dgt3 RNAi2</td><td>VDRC TRiP valium 20</td><td>BL58137</td><td>n/a</td></tr><tr><td>R68</td><td>dgt4 RNAi</td><td>VDRC KK</td><td>v108969</td><td>no</td></tr><tr><td>R63</td><td>dgt5 RNAi</td><td>VDRC TRiP valium 20</td><td>BL60366</td><td>n/a</td></tr></tbody></table>

<table><caption>Table S3</caption><thead><tr><th>Gene</th><th>Forward primer (5'-&gt;3')</th><th>Reverse primer (5'-&gt;3')</th></tr></thead><tbody><tr><td>a-tub84b</td><td>CACACCACCCTGGAGCATTC</td><td>CCAATCAGACGGTTCAGGTTG</td></tr><tr><td>SOP</td><td>CACCCCAATAAAGTTGATAGACCT</td><td>ACCACCACGAGAGCCAAAT</td></tr><tr><td>RPL32</td><td>CTTCATCCGCCACCAGTC</td><td>CGACGCACTCTGTTGTCG</td></tr><tr><td>GAPDH</td><td>GATGCTCCCATGTTCGTTTGC</td><td>GTCCATCACGCCACAACTTG</td></tr><tr><td>dgt4</td><td>CGCCATCATTGACAACTGAGG</td><td>GCTGTCTTCCTGAAAACGGC</td></tr><tr><td>grip91</td><td>TGTGGCAACCAATTCCACTGA</td><td>TTTTCGTGAACTTCGGGCATC</td></tr></tbody></table>
## Supplementary Movie Legends

**Movie S1. Movie showing EB1-GFP comets within the distal region of control-RNAi neurons.** Comets can be seen travelling towards the soma (retrograde) in the secondary dendrite and away from the soma in the terminal dendrite. Movie plays at 10 fps.

**Movie S2. Movie showing EB1-GFP comets within the distal region of Augmin-RNAi neurons.** Comets can be seen travelling both towards the soma (retrograde) and away from the soma (anterograde) in the secondary dendrite. Two terminal dendrites can be observed – an EB1-GFP comet travels within one terminal dendrite but no EB1-GFP comets travel in the other terminal dendrite. Movie plays at 10 fps.

**Movie S3. Movie showing terminal dendrite dynamics within the distal region of a control-RNAi neuron.** Distal region of a class IV da neuron expressing ppk-CD4-tdGFP and control-RNAi. Green arrows indicate terminal dendrites that are present at the beginning and the end of the movie. Red arrows indicate terminal dendrites that are present at the beginning of the movie but have disappeared by the end. Blue arrows indicate terminal dendrites that are absent at the beginning of the movie but appear during the movie. Movie plays at 5 fps.

**Movie S4. Movie showing terminal dendrite dynamics within the distal region of a Augmin-RNAi neuron.** Distal region of a class IV da neuron expressing ppk-CD4-tdGFP and Augmin-RNAi. Green arrows indicate terminal dendrites that are present at the beginning and the end of the movie. Red arrows indicate terminal dendrites that are present at the beginning of the movie but have disappeared by the end. Blue arrows indicate terminal dendrites that are absent at the beginning of the movie but appear during the movie. Movie plays at 5 fps.
## Supplementary Movie Legends

**Movie S1. Movie showing EB1-GFP comets within the distal region of control-RNAi neurons.** Comets can be seen travelling towards the soma (retrograde) in the secondary dendrite and away from the soma in the terminal dendrite. Movie plays at 10 fps.

**Movie S2. Movie showing EB1-GFP comets within the distal region of Augmin-RNAi neurons.** Comets can be seen travelling both towards the soma (retrograde) and away from the soma (anterograde) in the secondary dendrite. Two terminal dendrites can be observed – an EB1-GFP comet travels within one terminal dendrite but no EB1-GFP comets travel in the other terminal dendrite. Movie plays at 10 fps.

**Movie S3. Movie showing terminal dendrite dynamics within the distal region of a control-RNAi neuron.** Distal region of a class IV da neuron expressing ppk-CD4-tdGFP and control-RNAi. Green arrows indicate terminal dendrites that are present at the beginning and the end of the movie. Red arrows indicate terminal dendrites that are present at the beginning of the movie but have disappeared by the end. Blue arrows indicate terminal dendrites that are absent at the beginning of the movie but appear during the movie. Movie plays at 5 fps.

**Movie S4. Movie showing terminal dendrite dynamics within the distal region of a Augmin-RNAi neuron.** Distal region of a class IV da neuron expressing ppk-CD4-tdGFP and Augmin-RNAi. Green arrows indicate terminal dendrites that are present at the beginning and the end of the movie. Red arrows indicate terminal dendrites that are present at the beginning of the movie but have disappeared by the end. Blue arrows indicate terminal dendrites that are absent at the beginning of the movie but appear during the movie. Movie plays at 5 fps.