HOIL‐1 ubiquitin ligase activity targets unbranched glucosaccharides and is required to prevent polyglucosan accumulation

HOIL‐1 knockout (KO) mice exhibit early embryonic lethality (Fujita et al, 2018; Peltzer et al, 2018), but HOIL‐1 “KO” mice expressing low levels of a truncated protein comprising the N‐terminal ubiquitin‐like (UBL) and Npl4 zinc finger (NZF) domains, but lacking the C‐terminal region including the catalytic RBR domain (Fujita et al, 2018), accumulate polyglucosan in their brain, spinal cord and heart (MacDuff et al, 2015; Nitschke et al, 2022). Because HOIL‐1 stabilises its binding partner HOIP, the expression of HOIP and Sharpin is also greatly reduced in these HOIL‐1 “KO” mice. It was therefore unclear whether the accumulation of polyglucosan was caused by loss of the E3 ligase activity of HOIL‐1, loss of the E3 ligase activity of HOIP or reduced expression of the non‐catalytic domains of HOIL‐1, HOIP or Sharpin. In contrast, HOIL‐1, HOIP and Sharpin are expressed at normal levels in the E3 ligase‐inactive HOIL‐1[C458S] knock‐in mice that we have described previously (Kelsall et al, 2019b). We therefore investigated whether polyglucosan was present in the tissues of these mice. We found that polyglucosan did indeed accumulate in the brain of the HOIL‐1[C458S] mice, but not in their wild‐type littermates. Particularly high levels were present in the hind brain (pons) and the hippocampus, the extent of deposition being similar in mice varying in age from 0.5 to 1.5 years (Fig 1A–D). Similar to those found in human patients harbouring mutations in HOIL‐1 (Laforet et al, 2017), the polyglucosan deposits in the hippocampus and pons of HOIL‐1[C458S] mice were smaller and more speckled in appearance than those observed in other glycogenoses, such as those resulting from catastrophic general errors in glycogen synthesis, like the absence of glycogen branching enzyme.

Polyglucosan was present in lower amounts in the brain cortex (Fig EV1A and B). It was also present in smaller amounts in the heart, where it reached a maximum level after about a year (Fig 1E and F), and in the lungs and liver (Fig EV1C–F). These results establish that the E3 ligase activity of HOIL‐1 is required to prevent the accumulation of polyglucosan in several tissues.

We considered how HOIL‐1 might prevent the deposition of polyglucosan and wondered whether HOIL‐1 might not only ubiquitylate the hydroxyl side chains of serine and threonine residues in proteins but also the hydroxyl moieties present in glucose. To investigate whether HOIL‐1 was capable of ubiquitylating glycogen directly, we initially used an in vitro fluorescence‐based assay in which commercially available bovine liver glycogen was incubated with HOIL‐1, E1, E2, Mg²⁺‐ATP and fluorescent Cy5‐labelled ubiquitin. The conjugating enzyme UBE2L3 is strictly cysteine reactive and has been reported to be the physiologically relevant E2 conjugating for other RBR ligases, such as HHARI and HOIP (Lewis et al, 2015; Dove et al, 2017). We therefore included it as the E2 in our assays, as it is also likely to be the cognate E2 that pairs with HOIL‐1. Following SDS‐PAGE, the glycogen (detected by periodic acid‐Schiff [PAS] staining) is too large to enter the gel and remains at the origin (Fig 2A), as does a portion of the fluorescent Cy5‐labelled ubiquitin (Fig 2B, lanes 5 and 6). The comigration of Cy5‐labelled ubiquitin and glycogen required the presence of every reaction component. This high molecular mass Cy5‐ubiquitin signal was lost if the reaction was incubated with α‐amylase to degrade the glycogen (Fig 2B, lanes 7 and 8) or incubated with the oxyester‐specific nucleophile hydroxylamine (Fig 2B, lanes 9 and 10). These experiments indicated that HOIL‐1 had ubiquitylated glycogen and that ubiquitin was attached to it via an oxyester bond.

The majority of the glucose units in glycogen are linked via α‐1,4‐glycosidic bonds. The linear oligosaccharide maltoheptaose contains seven glucose units linked by such bonds (Fig 2C) and was used as a simple model substrate. The replacement of glycogen by maltoheptaose in the in vitro ubiquitylation reaction led to the formation of a single more slowly migrating form of ubiquitylated maltoheptaose in a concentration‐ and time‐dependent manner (Fig EV2A and B). As observed for glycogen, this adduct was sensitive to treatment with both hydroxylamine and α‐amylase, although the ubiquitin was still bound to maltose and maltotriose after digestion with α‐amylase because complete hydrolysis to glucose did not take place (Zakowski & Bruns, 1985) (Fig 2D, lanes 7 and 8). Mass spectrometry analysis of ubiquitylated maltoheptaose established that it was a monoubiquitylated species (Fig 2E).

To identify the ubiquitin ligation site, a hybrid labelled sample was prepared in which ubiquitin was labelled with NMR isotopes ¹⁵N and ¹³C and ligated to unlabelled maltoheptaose. The purpose was to reassign ubiquitin and measure NMR connectivity across the protein–carbohydrate interface. As anticipated, the ¹H‐¹⁵N HSQC spectrum of [¹⁵N,¹³C] ubiquitin maltoheptaose shows the characteristic amide peak patterns of ubiquitin and any peak movements could be reassigned readily. Two large amide chemical shift perturbations are observed for the C‐terminal residues G75 and G76, when compared to free ubiquitin (Fig 3A). Strikingly, the amide ¹⁵N and ¹HN signals of G76 are shifted upfield by −5.4 ppm and downfield by 0.3 ppm respectively. These values are consistent with those expected for C‐terminal amides when the charged carboxylate is removed by modification (Ulrich et al, 2008). No significant chemical shift perturbations are observed elsewhere in ubiquitin. Spin–spin relaxation time (T₂) of protein ¹⁵N NMR signals decreases with increasing molecular size. Measured T₂ values for ubiquitylated maltoheptaose lie in the 130–170 ms range, which agree with those measured for monomeric ubiquitin, compared to T₂ values of 90–100 ms for diubiquitin. Taken together, these data indicate that a single ubiquitin moiety is conjugated to the oligosaccharide via its C‐terminal glycine residue. Furthermore, the amide of G76 in ubiquitin maltoheptaose exhibits multiple chemical shift environments, with at least four resonances resolved (Fig 3A—boxed), which suggests that different isomers of the conjugate exist with individual maltoheptaose molecules being singly ubiquitylated at various sites along the oligosaccharide.

Comparison of the anomeric region of the 1D ¹H NMR spectrum for the ubiquitylated maltoheptaose with free maltoheptaose (Figs 3B and EV3A) shows that while the anomeric protons of the α and β reducing end sugar are largely unaffected, there are several significant chemical shift changes for the anomeric protons of the other glucose residues. As the anomeric position of these sugar residues cannot be targeted by the E3 ligase, the chemical shift changes reflect the effects of ubiquitylation on other positions within these sugars.

To identify unambiguously which position(s) on the glucose units were modified with ubiquitin, we used an ¹H‐¹³C heteronuclear multiple bond correlation (HMBC) experiment to detect the three‐bond scalar coupling between the sugar protons and the ¹³C carbonyl of G76. These ³J_COCH couplings are typically 2–3 Hz and correlations in the HMBC would reveal the specific ligated positions. Only one significant correlation could be observed at 4.30 and 4.50 ppm (Fig 3C), corresponding to two protons of a CH₂ group, and this can be assigned readily using ¹H correlation NMR spectroscopy to the side‐chain 6‐CH₂ position of the glucose residues. These chemical shifts match those for 6‐CH₂ groups typically found in (α1‐6) glucose chains (Dobruchowska et al, 2012), thereby confirming the 6‐position modification.

To provide further support for our assignment, we prepared a ubiquitylated maltose sample and compared the natural abundance ¹H‐¹³C HSQC spectrum with that for free ubiquitin (Figs 3D and EV3B). The 6‐CH₂ peaks at 4.30 and 4.50 ¹H ppm (67.0 ¹³C ppm) can be clearly observed in the ubiquitylated maltose sample and not ubiquitin. We also note a minor set of proximal CH₂ peaks in this region of the spectrum, which could either represent a different conformation of the non‐reducing 6‐CH₂ group due to restriction in free bond rotation from the attached ubiquitin or some ligation to the reducing end sugar 6‐CH₂ group, although there are no apparent chemical shift changes for the anomeric proton at the reducing end (Fig 3B). A second 6‐CH₂ group is also observed (3.85 and 3.90 ¹H ppm and 63.5 ¹³C ppm in Fig 3D), which corresponds to an unmodified glucose 6‐CH₂OH group (Bekiroglu et al, 2003) and this likely belongs to the reducing end carbohydrate residue of maltose. The ring CH protons are clearly visible between 71 and 81 ppm and these display no significant chemical shift changes when compared to free maltose, suggesting that any ubiquitylation of the ring CHOH groups is absent, or is minimal and beyond detection here.

Consistent with the NMR analysis, we found that cyclomaltoheptaose lacking the reducing C1 hydroxyl and non‐reducing C4 hydroxyl groups could still be ubiquitylated (Fig 3E and F), but the 6‐mercaptan derivative of cyclomaltoheptaose lacking any 6‐CH₂OH groups could not (Fig 3G and H). It should be noted that the hydroxymethyl (CH₂‐OH) group of a glycosyl unit is equivalent to the hydroxymethyl side chain of the amino acid serine, a previously identified target for HOIL‐1 ubiquitylation (Kelsall et al, 2019b).

Taken together, these experiments show that the single monoubiquitylated species of maltoheptaose observed on SDS gels is a mixture of products, each strictly monoubiquitylated on the C6 hydroxyl moiety but located on distinct single glycosyl units within the oligosaccharide. Since maltoheptaose forms a compact helical arrangement in solution (Goldsmith et al, 1982), steric considerations are likely to restrict ubiquitylation to a single site on maltoheptaose. The alternative explanation that ubiquitylation of the first glycosyl residue prevents the ubiquitylation of further glucosyl residues by disrupting the helical structure of maltoheptaose seems less likely, since cyclomaltoheptaose is also singly ubiquitylated, despite being restrained in a pseudohelical conformation by cyclisation that would not be disrupted when the first ubiquitin is added.

Since the rate of ubiquitylation of glycogen was slow, taking many hours to convert most of the ubiquitin in the assay to monoubiquitylated maltoheptaose (Fig EV2B), we wondered whether a mechanism(s) might be needed to accelerate the HOIL‐1‐catalysed ubiquitylation of glycogen in cells. We therefore initially investigated the binding of HOIL‐1 and other proteins to the unbranched α1:4‐linked polysaccharide amylose (Fig 4E) to evaluate their glucan‐binding properties (Fig 4A). Unlike maltose‐/maltodextrin‐binding protein (MBP), HOIL‐1 did not bind to amylose–agarose resin, similar to other negative control proteins, such as glutathione S‐transferase (GST). However, to our surprise, the other two components of LUBAC, Sharpin and HOIP did bind to amylose resin, the latter behaving similarly to MBP in the pull‐down assays (Fig 4A). To investigate whether the strength of interaction of HOIP and Sharpin with glucosaccharides depended on the extent of branching, we compared their ability to interact with amylopectin, a polysaccharide component of starch with few branch points, and highly branched glycogen from mammalian cells (Fig 4E). Strikingly, HOIP and Sharpin only bound to amylopectin in these assays (Fig 4B), indicating that these proteins bind preferentially to less branched polyglucosaccharides.

There are no obvious carbohydrate‐binding domains in the primary sequence of either HOIP or Sharpin. We therefore analysed various protein truncations for their ability to bind amylose in our pull‐down assay (Fig 4C and D). None of the truncated HOIP constructs bound to amylose, implying that non‐contiguous regions most likely contribute to glucan binding (Fig 4C). The isolated N‐terminal Pleckstrin homology (PH) domain of Sharpin did not bind to amylose resin, but removal of Sharpin’s Npl4 zinc finger (NZF) domain actually increased binding to amylose (Fig 4D). Constructs containing both the LUBAC‐tethering motif (LTM) and UBL domains of Sharpin interacted with amylose, whereas constructs lacking either of these domains did not (Fig 4D). This suggests that the LTM and UBL domains, known to play important roles in the stabilisation of the trimeric LUBAC (Fujita et al, 2018), may participate in binding to unbranched glucosaccharides.

The experiments described in the preceding section suggested a role for HOIP in recruiting HOIL‐1 to glycogen. We therefore wondered whether HOIP might also facilitate the HOIL‐1‐catalysed ubiquitylation of glycogen in other ways. HOIL‐1 contains an Npl4 zinc finger (NZF) domain that is reported to bind M1‐Ub dimers about 10‐fold more strongly than K63‐Ub dimers (Sato et al, 2011). We therefore wondered whether M1‐Ub oligomers, formed by the action of HOIP, might bind to the NZF domain of HOIL‐1, inducing a conformational change that accelerated the HOIL‐1‐catalysed ubiquitylation of maltoheptaose. We found that the inclusion of M1‐Ub or K63‐Ub dimers (Fig 5A) or tetramers (Fig 5B) did indeed speed up the rate of conversion of maltoheptaose to ubiquitylated maltoheptaose. Remarkably, after only 1 min, the conversion of ubiquitin to monoubiquitylated maltoheptaose was already greater than after 2 h in the absence of M1‐Ub or K63‐Ub oligomers. After 10–15 min, nearly all the ubiquitin in the assays had been converted to ubiquitylated maltoheptaose (Fig 5A and B). The experiment demonstrated that the rate of ubiquitylation of maltoheptaose was accelerated at least 100‐fold in the presence of small M1‐Ub or K63‐Ub oligomers.

The E2‐conjugating enzyme UBE2D3 (UbcH5c) has been reported to permit ester‐linked ubiquitylation when acting in combination with the E3 ligases MYCBP2 and RNF213 (Pao et al, 2018; Otten et al, 2021). We found that the rate of conversion of ubiquitin to ubiquitylated maltoheptaose was accelerated similarly by M1‐Ub or K63‐Ub oligomers when UBE2L3 was replaced by UBE2D3 (Fig EV4A), although the rate of ubiquitylation was much slower with UBE2D3 (compare Fig EV4A with Fig 5A and B). As expected, no ubiquitylation was observed in the assay with UBE2L3 when HOIL‐1 was replaced by the catalytically inactive HOIL‐1[C460A] mutant (Fig EV4B) (human C460 is equivalent to mouse C458).

To investigate whether the M1‐Ub and K63‐Ub oligomers exerted their effects on HOIL‐1 activity by binding to the NZF domain, we made mutations in this domain that have been reported to impair the binding of M1‐Ub dimers (Sato et al, 2011; Gomez‐Diaz et al, 2021). Surprisingly, these mutations did not reduce the rate of ubiquitylation of maltoheptaose by either M1‐Ub or K63‐Ub oligomers (Fig 5C). To check this result, we next made a mutant that removed most of the NZF domain, a deletion mutant that is known to abolish the binding of M1‐Ub dimers to the NZF domain (Sato et al, 2011). However, the activation of HOIL‐1 by M1‐Ub and K63‐Ub oligomers remained unimpaired (Fig 5D). Moreover, the N‐terminally truncated HOIL‐1[233‐510] mutant, which lacks the LTM, UBL and NZF domains, was also activated by M1‐Ub and K63‐Ub oligomers similarly to wild‐type HOIL‐1 (Fig 5E). Taken together, these experiments revealed that the small Ub oligomers did not activate HOIL‐1 by binding to the NZF domain or any of the other domains located in the N‐terminal region of the protein. They also suggested that HOIL‐1 contained another binding site(s) for M1‐Ub and K63‐Ub oligomers that was most likely located within the RBR domain itself.

The related RBR E3 ligase family members HOIP and Parkin are activated allosterically by the binding of M1‐Ub dimers (HOIP) and Ser65‐phosphorylated ubiquitin (Parkin) to a ubiquitin‐binding site within the RBR domain (Wauer et al, 2015; Lechtenberg et al, 2016). We therefore wondered whether the allosteric‐binding site for M1‐Ub dimers and K63‐Ub dimers in HOIL‐1 might be located in the analogous region of the protein and made mutations equivalent to those that impair activation of HOIP and Parkin by M1‐Ub dimers and phospho‐ubiquitin respectively (Fig 6A). Mutation of Ile358 to Arg (equivalent to the Ala320Arg mutation in Parkin that abolishes phospho‐Ub binding) or the double‐mutation Phe381Ala/Glu383Ala (equivalent to combining the Ile807Ala and Glu809Ala mutants that disrupt the allosteric activation of HOIP) resulted in a pronounced reduction or loss of HOIL‐1 activation by Ub dimers, but had less effect on the activation of HOIL‐1 by K63‐Ub tetramers and little effect on ligase activation by M1‐Ub tetramers (Fig 6B–D). We next combined these RBR domain mutations with the HOIL‐1[233‐510] truncation, which ablated the activation of HOIL‐1 by ubiquitin tetramers almost completely (Fig 6E and F). This suggested that, while not essential for the allosteric activation of HOIL‐1, a region(s) N‐terminal to the RBR may contribute to the binding of longer Ub oligomers.

In agreement with the data published using ubiquitin dimers (Sato et al, 2011; Gomez‐Diaz et al, 2021), we found that the mutation or loss of the NZF domain was sufficient to disrupt the binding of HOIL‐1 to immobilised Halo‐tagged M1‐Ub tetramers (Fig EV4C and D). This indicates that constructs containing the RBR domain, but lacking the NZF domain of HOIL‐1, have a much lower affinity for the M1‐Ub tetramer than constructs containing the NZF domain. This is in keeping with a model in which the binding of M1‐Ub oligomers to the NZF domain of HOIL‐1 anchors LUBAC at its sites of action, whereas the binding of M1‐Ub or K63‐Ub oligomers at the allosteric site on HOIL‐1, which activates this E3 ligase, is weaker and more transient.

Recently, DeepMind and EMBL’s European Bioinformatics Institute created the AlphaFold Protein Structure Database, making freely available structural predictions for all proteins in the human proteome (Jumper et al, 2021; Varadi et al, 2022). In order to gain more insight into how ubiquitin oligomers influence HOIL‐1 activity, we examined the HOIL‐1 structure predicted in the AlphaFold Protein Structure Database. Alignment of AlphaFold’s HOIL‐1 prediction with the experimentally derived structure of HOIP (Lechtenberg et al, 2016) revealed remarkable similarities in the conformations of the RING1 and in‐between‐RING (IBR) domains of these two proteins (Fig 6G), and suggested that the allosteric ubiquitin is accommodated by a common binding site in HOIL‐1 and HOIP (Fig 6G and H). Our mutagenesis data also revealed that hydrophobic interactions with ubiquitin’s Ile44 hydrophobic patch, coupled with the use of a glutamic acid “clamp” to bind the two arginines (Arg72/Arg74) at the C‐terminus of ubiquitin, are employed similarly in both proteins.

Interestingly, during the HOIL‐1‐catalysed reaction, the M1‐Ub or K63‐Ub dimers and tetramers included to accelerate the rate of formation of monoubiquitylated maltoheptaose, themselves became attached covalently to maltoheptaose and at a similar rate to the formation of monoubiquitylated maltoheptaose (Figs 5, 6, and EV4A and B). This en bloc transfer of pre‐assembled ubiquitin chains to maltoheptaose was dependent on UBE1 and UBE2L3, and did not require free monoubiquitin (Fig 7A and B). The oxyester bond linking maltoheptaose to the ubiquitin dimer or tetramer was hydroxylamine sensitive and could not be cleaved by the M1‐Ub‐specific deubiquitylase Otulin or the K63‐Ub‐specific deubiquitylase AMSH‐LP (Fig EV5A and B). Performing time‐course assays allowed the visualisation of a dithiothreitol‐sensitive tetraubiquitin adduct on UBE1, UBE2L3 and HOIL‐1 that was prominent at early time points but decreased after 2 min (Fig EV5C left panel). When wild‐type HOIL‐1 was replaced with a catalytically inactive C460S or C460A mutant, the transient adducts on UBE1 and UBE2L3 could not be discharged and persisted, while the use of the C460S mutant similarly stabilised a dithiothreitol‐insensitive oxyester‐linked tetraubiquitin adduct on HOIL‐1 (Fig EV5C, middle and right‐hand panels). These experiments are consistent with the HOIL‐1‐catalysed conjugation of ubiquitin oligomers to maltoheptaose en bloc. The HOIL‐1‐initiated polyubiquitylation of glycogen (and proteins) in cells might therefore be catalysed in a single step, and not by HOIL‐1‐catalysed monoubiquitylation followed by the sequential addition of further ubiquitin molecules through the action of other E3 ligases.

The finding that ubiquitin dimers and tetramers were substrates, as well as activators of HOIL‐1, raised the question of whether activation was dependent or independent of their ability to act as substrates. We therefore generated M1‐Ub dimers and tetramers in which the most distal C‐terminal Gly–Gly sequence was mutated to Ala–Ala. These mutants could no longer be conjugated to maltoheptaose, but remained equally potent activators of the HOIL‐1‐catalysed monoubiquitylation of maltoheptaose (Fig 7C). The rate of ubiquitylation of maltoheptaose did not decrease significantly if the M1‐Ub tetramer concentration in the assay was decreased from 5.0 to 0.5 µM at a constant HOIL‐1 concentration of 2.0 µM (Fig 7D), indicating that they were acting catalytically. Taken together, the results support the conclusion that ubiquitin oligomers activate HOIL‐1 allosterically and not by acting as alternative substrates of this E3 ligase.

E3 ligase‐inactive HOIL‐1[C458S] mice (Kelsall et al, 2019b) and wild‐type littermates were euthanised by injecting a lethal dose of sodium pentobarbitone. The brain, heart, lungs and liver were removed and fixed for 48–72 h in 10% neutral buffered formalin. Tissues were processed and 3 μm deparaffinised sections were treated for 45 min with 0.5% (w/v) α‐amylase (Sigma), rinsed for 30 min in water and processed for periodic acid‐Schiff (PAS) staining. The tissue sections were assessed by a veterinary pathologist (C.S.) blinded to the genotype of the mice in the different cohorts. A simple semi‐quantitative scoring system was used for the assessment of α‐amylase‐resistant PAS‐positive materials. Scores for amylase‐resistant PAS‐positive deposits were defined as: (i) occasional granular deposits, just detectable by light microscopy; (ii) increased numbers of deposits, readily detectable at ×10 magnification, and (iii) markedly increased numbers of granular deposits easily identifiable by light microscopy. The photomicrographs were captured with Nanozoomer software from whole slide image scans prepared using a Hamamatsu slide scanner.

DNA constructs were generated by MRC‐PPU Reagents & Services and sequenced by MRC‐PPU DNA Sequencing & Services (www.dnaseq.co.uk). All plasmids used in this study are available upon request at https://mrcppureagents.dundee.ac.uk. Full‐length untagged human HOIL‐1 (and various mutants thereof) was expressed as His₆‐SUMO‐HOIL‐1 in BL21 (DE3) E. coli cells before removal of the tag with the SUMO protease Ulp1 and buffer exchange by size exclusion chromatography or dialysis into PBS, 1 mM TCEP. Human ubiquitin, GST‐Otulin and GST‐AMSH‐LP [246‐436] were expressed and purified as described elsewhere (Ritorto et al, 2014). The expression and purification of His₆‐UBE1 and UBE2L3 (Kelsall et al, 2019b), UBE2D3 (Pao et al, 2018), GST‐CHIP (Zhang et al, 2005) and HHARI (Duda et al, 2013) have been described previously. HOIP and Sharpin were expressed in BL21 cells as GST‐tagged fusion proteins, purified on GSH Sepharose and collected by elution with 10 mM glutathione or by removal of the GST tag on the resin using PreScission Protease. MBP‐His₆ was purified by Ni²⁺ Sepharose chromatography. Untagged Lys63‐linked and linear di‐ and tetraubiquitin chains were produced and purified according to previously reported methods (Komander et al, 2008; Dong et al, 2011). His‐Halo‐tagged M1‐Ub tetramers were purified by Ni²⁺ Sepharose chromatography and ubiquitin oligomers in which the C‐terminal Gly–Gly sequence of the distal ubiquitin was mutated to Ala–Ala were purified by means of an N‐terminal, TEV protease‐cleavable, DAC tag (Lee et al, 2012). All proteins were prepared in PBS, 1 mM TCEP and stored in aliquots at −80°C.

Reactions (20 μl) contained 500 nM His₆‐UBE1, 2 μM UBE2L3, 2 μM HOIL‐1, 10 μM Cy5‐ubiquitin (South Bay Bio) and 10 mg/ml bovine liver glycogen (Sigma) in phosphate buffered saline pH 7.4 containing 0.5 mM TCEP and 5 mM Mg²⁺‐ATP. Reactions were incubated at 37°C for 1 h with gentle mixing. Where indicated, reactions were then incubated in the presence of 50 μg/ml (~0.1 units) human salivary α‐amylase (Sigma) or 1.5 M hydroxylamine for a further 60 min at 37°C. Reactions were stopped by adding NuPAGE LDS sample loading buffer supplemented with 50 mM DTT, and denatured at room temperature for 10 min. Reaction products were separated by gel electrophoresis on 4–12% Bis–Tris gradient gels using MES SDS running buffer (Invitrogen) and stained with Flamingo fluorescent protein stain (Bio‐Rad). Visualisation of the fluorescent signals was performed using the ChemiDoc MP Imaging System (Bio‐Rad) and ImageJ software (Schneider et al, 2012). In the case of assays using glycogen as substrate, these gels were then stained with periodic acid‐Schiff stain using the Pierce Glycoprotein Staining Kit (Thermo Scientific) and Coomassie stained with InstantBlue Protein Stain (Expedeon).

Reactions (typically 10–20 μl) contained 500 nM His₆‐UBE1, 2 μM UBE2L3, 2 μM HOIL‐1, 10 μM ubiquitin and 20 mM maltosaccharide substrate (unless otherwise indicated) in phosphate buffered saline pH 7.4, 0.5 mM TCEP and 5 mM Mg²⁺‐ATP. Reactions were incubated at 37°C for 1 h with gentle mixing, unless otherwise stated. Where indicated, reactions were then incubated in the presence of 50 μg/ml (~0.1 units) human salivary α‐amylase or 1.5 M hydroxylamine for a further 60 min at 37°C. Reactions were quenched by addition of NuPAGE LDS sample loading buffer supplemented with 50 mM DTT, and denatured at room temperature for 10 min. Reaction products were separated by gel electrophoresis on 4–12% Bis–Tris gradient gels using MES SDS running buffer and stained with Coomassie InstantBlue Protein Stain before analysis on the ChemiDoc MP Imaging System. Maltoheptaose was purchased from CarboSynth. Maltose was from Sigma. Cyclomaltoheptaose (β‐cyclodextrin) and 6‐mercapto‐cyclomaltoheptaose (heptakis‐(6‐deoxy‐6‐mercapto)‐β‐cyclodextrin) were from AraChem. The UBE1 inhibitor MLN7243 was purchased from Active Biochem.

Where noted, HOIL‐1 activity was stimulated by addition of chain‐type‐specific ubiquitin dimers or tetramers at a concentration of 2 μM. In these experiments, the ubiquitin chains were pre‐incubated for 30 min at 30°C with all other reaction components except ATP. Reactions were initiated by adding Mg²⁺‐ATP to a final concentration of 5 mM. In order to slow the reaction rate and allow better visualisation of the differences between ligase activation in the presence of different chain types, maltoheptaose concentration was dropped to 10 mM and reactions proceeded at the lower temperature of 30°C for the indicated times.

Ubiquitylation reactions were carried out as described previously and terminated through the addition of 20% (v/v) trifluoroacetic acid (TFA) to a final concentration of 2%. Ubiquitylated maltoheptaose was separated from unconjugated free ubiquitin by reverse‐phase–high‐performance liquid chromatography (RP–HPLC) using a Dionex Ultimate 3000 System. A Thermo BioBasic column (250 × 10 mm) was equilibrated in aqueous buffer containing 20% (v/v) acetonitrile supplemented with 0.1% (v/v) TFA. A flow rate of 2.3 ml/min and an increasing gradient of acetonitrile from 20% to 60% over 60 min were utilised for sufficient separation. Separated fractions were validated using MALDI‐TOF mass spectrometry before being pooled and freeze‐dried. Ubiquitylated maltose was purified similarly.

For MALDI‐TOF sample preparation, a mixture containing a 1:1 ratio of 2% (v/v) TFA and 2,5 dihydroxyacetophenone (DHAP) matrix solution (7.6 mg of 2,5 DHAP in 375 μl 100% (v/v) ethanol and 12 μl of 12 mg/ml diammonium hydrogen citrate) was added to the sample in a 1:1 ratio. One microlitre of the solutions were spotted in duplicate onto an MTP AnchorChip 1,536 Target (Bruker Daltronics). Samples were air dried at room temperature prior to analysis. All spectra were acquired using a Rapiflex MALDI‐TOF mass spectrometer (Bruker Daltronics) equipped with Compass 1.3 control. Recording took place in automatic mode (AutoXecute, Bruker Daltronics), allowing 6–9 s per target spot as described previously (De Cesare et al, 2020). Spectra were visualised using FlexControl software and processed by FlexAnalysis software (version 4.0).

[U‐¹⁵N,¹³C]‐labelled Ubiquitin was produced by growing transformed E. coli BL21 (DE3) cells (New England Biolabs) in M9‐enriched medium supplemented with ¹⁵NH₄Cl (1 g/l) and ¹³C‐glucose (2 g/l). All spectra were recorded on a Bruker Avance III HD 800 MHz equipped with triple‐resonance cryoprobe. Purified [U‐¹⁵N,¹³C]‐labelled ubiquitin maltoheptaose conjugate was freeze dried and dissolved in 0.5 ml of 20 mM phosphate buffer pH 6.5, 150 mM NaCl and 10% D₂O to a final concentration of 0.5 mM. The sequential NMR assignment of ubiquitin was amended using HN(CO)CACB; HNCACB, HN(CA)CO and HNCO triple‐resonance spectra were acquired at 303 K (Cavanagh et al, 2007). 2D heteronuclear spectra such as ¹H‐¹³C heteronuclear multiple bond correlations (HMBC) spectroscopy and heteronuclear single quantum correlation (HSQC) were recorded with standard experiments.

Amylose covalently bound to agarose resin (New England Biolabs) was pre‐incubated with 10 mg/ml bovine serum albumin (BSA) for 30 min with mixing at room temperature to block non‐specific binding. The resin was pelleted by centrifugation (500 g for 2 min) and washed three times with 10 mM phosphate buffer, pH 7.4, containing 137 mM NaCl, 1 mM TCEP, 0.01% (v/v) Tween‐20 and 0.01 mg/ml BSA (binding buffer). One microgram of bacterially expressed protein was then incubated with 30 μl of this blocked amylose resin with mixing for 60 min at 4°C in a final volume of 150 μl of binding buffer. Following incubation, the beads were pelleted by centrifugation (500 g for 2 min), the supernatant removed and retained, and the resin washed three times with binding buffer before finally resuspending with buffer to the original 150 μl starting volume. Bound proteins were eluted by addition of NuPAGE LDS gel loading buffer and proteins in the supernatant and pellet were analysed by gel electrophoresis followed by immunoblotting using the following antibodies: anti‐HOIP (#SAB2102031) and anti‐HOIL‐1 (#HPA024185) were from Sigma‐Aldrich; anti‐MBP (#E8032S) was from New England Biolabs; HRP‐conjugated anti‐rabbit secondary (#7074) and HRP‐conjugated anti‐mouse secondary (#7076) were from Cell Signaling Technology and HRP‐conjugated anti‐sheep secondary (ab97130) was from Abcam. Polyclonal antibodies against GST (#S902A), CHIP (#S471B) and HHARI (#S622D) were generated by MRC‐PPU Reagents & Services and are available upon request at https://mrcppureagents.dundee.ac.uk.

The binding of bacterially expressed proteins to bovine liver glycogen (Sigma) and potato starch amylopectin (Sigma) was examined as described (Jiang et al, 2010). GST‐HOIP, GST‐Sharpin (1‐308) or untagged HOIL‐1 were diluted in 50 mM Tris–HCl, pH 7.5, 150 mM NaCl, 0.1% (v/v) 2‐mercaptoethanol, 0.1% (v/v) Triton X‐100 and subjected to centrifugation at 10,000 g for 20 min at 4°C. The supernatant was mixed with the supernatant of glycogen/amylopectin centrifuged under the same conditions. The final concentration of protein was 2.5 μg/ml and glycogen/amylopectin was 0.25 mg/ml in a volume of 0.5 ml. Protein in the presence of buffer alone was used as a negative control. The samples were incubated for 1 h at 4°C with end‐over‐end mixing, and then subjected to ultracentrifugation for 90 min at 100,000 g at 4°C. The pellets were resuspended in buffer to the starting volume of 0.5 mL and proteins in the supernatant and pellet were visualised by immunoblotting with the antibodies indicated.

HOIL‐1 binding to Met1‐linked tetraubiquitin was performed using a modification of previously described methods (Kelsall et al, 2019a). 0.1 mg of Halo‐tagged tetraubiquitin was coupled to 0.1 ml of HaloLink resin (Promega) in 1 ml of coupling buffer (phosphate buffered saline pH 7.4, 0.001% NP‐40 and 1 mM TCEP) overnight at 4°C. The beads were washed five times with binding buffer (phosphate buffered saline pH 7.4, 0.001% NP‐40, 1 mM TCEP and 0.01 mg/ml BSA), then 10 μl of coupled beads was incubated with 2 μg of bacterially expressed HOIL‐1 protein in 0.5 ml of binding buffer for 3.5 h at 4°C. The beads were washed five times with phosphate buffered saline pH 7.4, 0.001% NP‐40 and 1 mM TCEP. Bound proteins were separated on 4–12% Bis–Tris gels and Coomassie stained with InstantBlue Protein Stain.