ER nests are specialized ER subdomains in Arabidopsis where peroxisomes and lipid droplets form

Zachary J Wright; Nathan E Tharp; Bonnie Bartel

doi:10.1016/j.devcel.2025.03.005

. Author manuscript; available in PMC: 2026 Mar 28.

Published in final edited form as: Dev Cell. 2025 Mar 28;60(15):2061–2080.e4. doi: 10.1016/j.devcel.2025.03.005

ER nests are specialized ER subdomains in Arabidopsis where peroxisomes and lipid droplets form

Zachary J Wright ^1,², Nathan E Tharp ¹, Bonnie Bartel ^1,^3,^*

PMCID: PMC12324974 NIHMSID: NIHMS2066114 PMID: 40157364

SUMMARY

Organelles are defining features of eukaryotic cells, yet much remains to be learned about organelle biogenesis. Lipid droplets and peroxisomes, which play opposing roles in storing and catabolizing fats, form from a mysterious domain in the endoplasmic reticulum (ER). We used live-cell fluorescence microscopy to visualize peroxisome and lipid droplet biogenesis in young Arabidopsis seedlings, where lipid catabolism is active and peroxisomes can be unusually large. We found that the ER domains where these organelles are born, which we term ER nests, are complex, dynamic structures that exclude general ER proteins but accumulate other proteins, including lipid biosynthetic enzymes and the COPII component SAR1. Furthermore, ER nests appear to define peroxisome-lipid droplet contact sites. Our findings provide a framework for understanding how these domains form and sort their protein components, illuminate eukaryotic lipid biosynthesis, and elucidate how distinct organelles arise from the ER.

Graphical Abstract

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eTOC blurb

Wright et al. used live-cell fluorescence microscopy of Arabidopsis seedlings stably transformed with various reporter constructs to reveal discrete ER domains (ER nests) with diverse cellular functions, including hosting peroxisome and lipid droplet biogenesis, compartmentalizing lipid synthesis, and influencing organelle contact sites.

INTRODUCTION

A central question of cell biology is how organelles of the endomembrane system, including peroxisomes and lipid droplets (LDs), acquire distinct identities despite originating from an entirely different organelle, the endoplasmic reticulum (ER). Peroxisomes are essential for life in most multicellular eukaryotes, including humans¹ and plants². Peroxisomes are often thought to be simple organelles in which a single lipid bilayer encloses a dense protein-only lumen; however, we recently documented additional complexity in the form of extensive intralumenal vesicles (ILVs) within plant peroxisomes³. Peroxisomes can form via division or by budding from the ER in an enigmatic de novo biogenesis process⁴. Lumenal proteins are imported into peroxisomes after cytosolic receptors recognize peroxisome targeting signals (PTSs), usually a C-terminal PTS1⁵. Peroxisomal membrane proteins are recognized via a membrane PTS (mPTS) and either directly inserted into peroxisome membranes or indirectly targeted to nascent peroxisomes from the ER⁶. The proteins needed to build peroxisomes are known as peroxins (PEX proteins).

Peroxisomes compartmentalize various oxidative reactions, usually including fatty acid β-oxidation, the earliest traceable peroxisomal pathway among eukaryotes⁷. Before β-oxidation, fats are stored in LDs, which are delimited by a phospholipid monolayer and enclose a core of triacylglycerols (TAGs) and sterol esters⁸. Like peroxisomes, LDs arise from the ER⁹, a multifunctional organelle that can display tubular, sheet-like¹⁰, or more complicated morphologies. The nuclear envelope is specialized ER that compartmentalizes the genetic contents of eukaryotic cells, and ER exit sites orchestrate cargo packaging and budding for delivery to other compartments. The ER influences other organelles, defining mitochondrial¹¹ and endosomal¹² fission events. Interestingly, peroxisomes and LDs appear to form at the same ER subdomains in yeast and mammalian cells¹³, and the biogenesis of both organelles is coregulated in C. elegans¹⁴. Much remains unknown about how ER domains are partitioned, the functions of specific subdomains, how the ER controls other organelles, and how distinct organelles arise from the ER.

Dysfunctional peroxisomes and LDs underlie various diseases^1,15 and are implicated in cancer^16,17 and aging^18,19. Investigating peroxisome and LD biogenesis from the ER has been limited by technical challenges in observing these structures as they form, particularly because their sizes in yeast and mammalian cells are near or below the resolution of light. Arabidopsis is well-suited to interrogating peroxisomes² and LDs²⁰ and can have strikingly large cells and organelles, allowing visualization of peroxisomes with remarkable sub-organellar detail in living plants³.

In this work, we gained unprecedented insight into the structure of the ER subdomain where peroxisomes and LDs are born by generating a suite of Arabidopsis lines stably transformed with various constitutive and inducible fluorescent reporters and examining these reporters using live-cell and time-lapse confocal microscopy. We observed these organelles forming within specific ER domains—ER nests—that exclude general ER proteins but concentrate biogenesis-related proteins, lipid biosynthetic enzymes, and SAR1 isoforms. These apparent non-canonical ER exit sites are shaped by atlastin and can remain in contact with their progeny organelles. Moreover, these dynamic domains can move back to and surround peroxisomes during conditions of increased metabolic activity.

RESULTS

De novo peroxisome formation is active and can be observed in Arabidopsis seedlings

De novo peroxisome formation has not been observed in a non-mutant context, limiting understanding of peroxisome biogenesis. To investigate whether de novo peroxisome formation occurs in Arabidopsis, we sought to identify pre-peroxisomes—immature peroxisomal membranes that had not yet imported lumenal proteins—using a line³ expressing a red peroxisome lumenal reporter (mRuby3-PTS1) and a green peroxisome membrane reporter targeted to peroxisomes via the C-terminal region of PEX26 [mNeonGreen (mNG)-mPTS^PEX26] (Figure 1A). We used live-cell confocal microscopy to image young seedlings, where abundant peroxisomes are catabolizing the seed LDs that fuel growth before photosynthesis is established. We observed both mature peroxisomes, defined by co-localizing membrane and lumenal reporters, and plentiful pre-peroxisomes—vesicles marked only by the membrane reporter (Figure 1B). Time-lapse imaging showed lumenal reporter import into the vesicles (Figure 1C), confirming that they were indeed pre-peroxisomes and establishing that de novo peroxisome formation naturally occurs in Arabidopsis.

Figure 1. — (A) Classic model of peroxisome formation from the ER. Peroxisomal membrane proteins sort to ER subdomains that bud pre-peroxisomes, which mature and acquire lumenal protein and ILVs.

(B) Pre-peroxisomes visualized in Arabidopsis seedlings as mNeonGreen (mNG)-mPTS^PEX26-positive vesicles lacking lumenal mRuby3-PTS1 (arrows).

(C) Time-lapse microscopy of panel B reporter showing mNG-mPTS^PEX26-positive vesicles (arrows) acquiring lumenal protein.

(D) In 6-day-old seedlings after 1 day of mPTS^PEX22-mTagBFP2 induction, mPTS^PEX22-mTagBFP2 localizes to ER but not pre-existing peroxisomes (mNG-mPTS^PEX26 and tdTomato-PTS1, arrows, top), whereas in 6-day-old seedlings after 6 days of induction, mPTS^PEX22-mTagBFP2 also localizes to peroxisomes (arrows, bottom). Line scans show relative intensities of mPTS^PEX22-mTagBFP2 (blue), mNG-mPTS^PEX26 (green), and tdTomato-PTS1 (magenta) of selected peroxisomes (red arrow in merge).

(E) mPTSP^EX22-mTagBFP2 labels the general ER but concentrates in puncta (arrowheads) that do not accumulate the general ER reporter (mNG-CB5DTMD). mPTS^PEX22-mTagBFP2 also labels some sheets (asterisks) that exclude the general ER reporter.

(F) Peroxisome (TdTomato-PTS1, arrow) enclosed in a structure with concentrated mPTS^PEX22-mTagBFP2 and excluding mNG-CB5DTMD (arrowhead). Line scans show the relative intensities of mPTS^PEX22-mTagBFP2 (blue), mNG-CB5DTMD (green), and tdTomato-PTS1 (magenta) of a peroxisome in an ER nest (red arrow in merge).

(G) The mNG-mPTS^PEX26 reporter labels peroxisomes (arrows) appearing to form within mPTS^PEX22-mTagBFP2-positive structures (arrowheads). Line scans show the relative intensities of mPTS^PEX22-mTagBFP2 (blue), mNG-mPTS^PEX26 (green), and tdTomato-PTS1 (magenta) of an ER nest (red arrow in merge).

(H, I) mPTS^PEX22-mNG-positive domains enclosing peroxisomes (mRuby3-PTS1; arrows) are present in the autophagy null mutant *atg7-4*, demonstrating that they are not autophagosomes.

(J) Hypothetical model of peroxisome formation in ER nests. A subset of peroxisomal membrane proteins sort to and concentrate in ER subdomains (ER nests), and peroxisomes bud from the inner layer.

Although pre-peroxisomes presumably bud from the ER, we did not observe notable ER localization of the mNG-mPTS^PEX26 peroxisome membrane reporter. However, we previously observed both ER and peroxisomes using a peroxisome membrane reporter targeted by the N-terminal region of PEX22 (mPTS^PEX22)³. To determine whether this localization reflected direct sorting to both membranes or trafficking from the ER to peroxisomes during de novo formation, we generated an Arabidopsis line inducibly expressing mPTS^PEX22-mTagBFP2 and constitutively expressing peroxisomal membrane (mNG-mPTS^PEX26) and lumenal [tdTomato (tdT)-PTS1] reporters (Figure S1). Like our previous PEX22-based reporter³, mPTS^PEX22-mTagBFP2 sorted to both ER and peroxisomes (Figure S2B). To explore the timing of this localization, we examined 6-day-old seedlings following either overnight (pulsed) or 6-day (continuous) reporter induction. Plants with pulsed mPTS^PEX22 expression showed reporter in the ER but not peroxisomal membranes, whereas seedlings of the same age with continuous expression showed both ER and peroxisome membrane localization, including ILVs (Figure 1D). These distinct localizations suggest that PEX22 traffics to peroxisomes via the ER during de novo formation rather than inserting directly into peroxisome membranes (Figure 1A).

De novo peroxisome formation occurs at discrete ER domains—ER nests

Peroxisome formation from the ER presumably requires a distinct domain to accumulate membrane proteins destined for peroxisomes while excluding general ER membrane proteins. Because the mPTS^PEX22 reporter appeared to traffic through the ER, we reasoned that the reporter might accumulate in such ER domains. We therefore combined our mPTS^PEX22 reporter with a general ER membrane reporter—a fusion of mNeonGreen to the transmembrane domain (TMD) of the ER-resident cytochrome B5-D (CB5D), which localizes fluorescent proteins to the Arabidopsis ER membrane²¹. We constitutively expressed mNG-CB5DTMD to label ER, mPTS^PEX22-mTagBFP2 to label ER and peroxisome membranes, and tdT-PTS1 to label peroxisomal lumen. As expected, mNG-CB5DTMD labeled ER but not peroxisomal membranes (Figure S2C). The mPTS^PEX22 reporter localized to peroxisomes and ER, including some ER sheets where the mNG-CB5DTMD reporter did not concentrate (Figure 1E). In young seedlings, where we observed abundant pre-peroxisomes (Figure 1B), the mPTS^PEX22 reporter concentrated in domains that largely excluded the mNG-CB5DTMD general ER reporter (Figure 1F, S2C, S2E) but contained an ER lumenal reporter (Figure S2D), suggesting that these regions were specialized ER-sorting domains. These domains did not accumulate mNGmPTS^PEX26, suggesting that they were not peroxisomal membranes, yet they frequently enclosed mNG-mPTS^PEX26 and/or tdT-PTS1-marked peroxisomes (Figure 1F, 1G). Interestingly, these domains often appeared thicker than a single lipid bilayer (Figure 1G, S2C), hinting at a multilamellar structure. Moreover, enclosed peroxisomes appeared to be budding from the inner membrane layer (Figure 1G).

These mPTS^PEX22 reporter-marked structures were distinct from previously characterized compartments, including endosomes (Figure S2F) and vacuoles (Figure S2E), the plant lysosome equivalent. Moreover, we confirmed that these structures were not autophagosomes by visualizing these structures enclosing peroxisomes in the atg7-4 null mutant²², in which autophagy is prevented (Figure 1H, 1I). We refer to these specialized ER domains giving rise to peroxisomes as “ER nests” (Figure 1J).

Lipid droplets can form from ER nests

Because LDs form at the same ER subdomains as peroxisomes in yeast and mammalian cells¹³, we examined our reporters during LD biogenesis. Arabidopsis LDs are primarily formed during embryogenesis to store fixed carbon for peroxisomal catabolism in seedlings before photosynthesis is established. To circumvent the technical difficulty of observing embryonic LD biogenesis, we incubated seedlings with oleate, a fatty acid that induces LD formation in yeast and mammalian cells²³.

Whereas mock-treated 6-day-old seedlings had few remaining seed LDs, we observed many large LDs after overnight oleate (0.05%) treatment of 5-day-old seedlings (Figure 2A, 2B). These LDs were visible using fluorescent dyes that accumulate in neutral lipids and via altered diffraction in bright-field images (Figure 2A). Unlike in yeast, which proliferate peroxisomes in response to oleate²³, peroxisome numbers did not significantly change following oleate treatment (Figure 2A, 2B).

We readily observed LDs enclosed in mPTS^PEX22-positive membranes following oleate treatment (Figure 2C), suggesting that LDs, like peroxisomes, form in ER nests. Moreover, we observed oleate-induced LDs enveloped in ER using an ER lumenal reporter²⁴ (Figure S2G–I), confirming that these structures were indeed ER-derived and not ER membrane reporter artifacts.

We investigated the relationship between ER nests and LD biogenesis by localizing two proteins involved in LD formation: diacylglycerol acyltransferase (DGAT) and seipin. DGAT is the final enzyme in TAG synthesis and localizes to LDs and the ER domains where LDs originate²⁵; DGAT overexpression increases LD formation in plants²⁶. Seipin specifies LD biogenesis sites on the ER²⁷ and localizes to puncta on the ER or around LDs²⁸, often at ER-LD junctions^28,29. We generated lines inducibly expressing DGAT1 or SEIPIN3 fused to mTagBFP2 and constitutively expressing mPTS^PEX22-mNG and tdT-PTS1 (Figure S1). After DGAT1 induction, we observed LDs enclosed within DGAT1- and mPTS^PEX22-labeled nests in young seedlings (Figure 2D), again suggesting that ER nests house LD biogenesis. Similarly, after co-incubating 5-day-old seedlings with estradiol and oleate overnight to induce seipin expression and LD formation, we observed seipin reporter in mPTS^PEX22-labeled ER nests around peroxisomes and LDs, sometimes within the same nest (Figure 2E). LDs budding out of ER nests were enclosed in residual seipin- and mPTS^PEX22-labeled membrane (Figure 2E, top row), and some seipin-positive ER nests contained internal nest membranes near peroxisomes marked with the mPTS^PEX22 reporter but without seipin (Figure 2E, bottom row), suggesting sorting within nests to exclude LD proteins from forming peroxisomes.

Because the ER nests we observed using fluorescence microscopy often appeared morphologically complex and potentially thicker than a single membrane, we sought to examine ER nest morphology using transmission electron microscopy (TEM) on wild-type seedlings lacking reporters. We could not definitively identify ER nests or most peroxisomes in younger seedlings (perhaps due to poor contrast of peroxisome and ER membranes in young tissue). After oleate application, however, we observed multilamellar structures that appeared to be smooth ER enclosing cytosol (Figure 2F) or LDs (Figure 2G). We concluded that LDs, like peroxisomes, can form from specialized ER subdomains—ER nests—in Arabidopsis (Figure 2H).

Lumenal protein import drives peroxisome expansion in ER nests

We often observed that peroxisomes appearing to bud from the innermost nest membrane already contained lumenal proteins (Figure 1F–I). We hypothesized that de novo peroxisome formation might involve membrane peroxins sorting from the cytosol into the inner layer of ER nests, allowing nascent pre-peroxisomes to enlarge via lumenal protein import. We tested this hypothesis by examining our reporters in the pex5-10 mutant³⁰, which has impaired peroxisomal lumenal protein import stemming from reduced function of the PTS1 receptor, PEX5^31,32. LDs were abundant in 2-day-old wild-type seedlings, and peroxisomes were present as distinct organelles with lumenal protein and ILVs (Figure 3A, 3B). ER nests were present as mPTS^PEX22-labeled structures that regularly enclosed LDs but did not concentrate mNG-mPTS^PEX26 (Figure 3A). In older seedlings, we observed some mPTS^PEX26 reporter in small ER nests dispersed in the general ER (Figure 3E), suggesting a small amount of mPTS^PEX26 reporter may fail to enter budding peroxisomes in wild-type cells. In contrast, young pex5-10 seedlings had few, if any, peroxisomes, as the lumenal reporter was largely dispersed in the cytosol (Figure 3C), and the mNG-mPTS^PEX26 reporter was often concentrated in ER nests enclosing LDs (Figure 3C, 3D). In older pex5-10 seedlings, we observed strong mPTS^PEX26 localization in small ER nests and some localization in the general ER (Figure 3F). The accumulation of peroxisomal membrane reporter in ER nest membranes when lumenal protein import is impaired implies that lumenal protein import facilitates nascent peroxisome budding from ER nests (Figure 3H) and suggests a distinct biogenesis mechanism from that used in canonical ER exit site budding (Figure 3G).

Figure 3. — (A) In young wild-type seedlings, the peroxisomal membrane reporter mNG-mPTS^PEX26 concentrates on peroxisomes (tdT-PTS1; arrow) but not ER nests (mPTS^PEX22-mTagBFP2-positive, mNG-mPTS^PEX26-negative structures), which often enclose LDs (LipidTOX Deep Red; yellow; arrowhead). Individual channels at right are single slices from boxed area.

(B) Section of panel A image showing mNG-mPTS^PEX26 (green) not localizing around LDs (magenta) in wild-type seedlings.

(C) In young *pex5-10* seedlings, tdT-PTS1-positive peroxisomes are not visible, and mNG-mPTS^PEX26 concentrates with mPTS^PEX22-mTagBFP2 in ER nests (arrowhead) that often enclose LDs. Individual channels at right are single slices from boxed area.

(D) Section of panel C image showing mNG-mPTS^PEX26 (green) localizing in ER nests (arrowheads) around LDs (magenta) in *pex5-10* seedlings.

(E, F) In 8-day-old seedlings, mNG-mPTS^PEX26 (green) localizes strongly to peroxisomes (magenta, arrow) and faintly to ER nests (blue, arrowheads) in wild type (E) but accumulates in ER nests (arrowheads) and the general ER in *pex5-10* (F).

(G) In classical ER exit sites, coat proteins corral lumenal cargo into budding vesicles to ferry ER lumen-derived cargo, which requires lumenal connectivity of the exit site with the general ER.

(H) ER nests are expected to limit lumenal connectivity to exclude general ER proteins and concentrate biogenesis-related proteins. Nascent peroxisome expansion may be driven via peroxisome lumenal protein import from the cytosol.

Sterol biosynthetic enzymes localize to ER nests, and nest membrane remains with mature lipid droplets

Because LDs contain sterol esters in addition to TAG, we investigated whether nests harbored sterol biosynthetic enzymes (Figure 4A). HMG-CoA reductase (HMGR) is a multi-pass membrane protein that in yeast localizes to specific ER domains that can enclose LDs³³. We observed co-localization of an inducible HMGR1-mTagBFP2 reporter with mPTS^PEX22-mNG–labeled ER nests, often around LDs or peroxisomes (Figure 4B–4D). In contrast, we did not observe HMGR1 on peroxisome membranes (Figure 4B–4D), suggesting exclusion from budding peroxisomes, similar to our seipin reporter (Figure 2E).

We also examined the HMGR1 reporter in older seedlings and after inducing LD biogenesis with oleate. We incubated 4-day-old seedlings with estradiol overnight to induce reporter expression, then transferred the seedlings to media without estradiol and with or without oleate for 1 day (Figure S3A). In mock-treated seedlings, the HMGR1 reporter localized to discrete puncta that overlapped with the mPTS^PEX22 nest reporter (Figure 4C) but minimally co-localized with mature peroxisomes (Figure 4C, S3D). In oleate-treated seedlings, the HMGR1 reporter mostly localized to ER membrane around newly-formed LDs (Figure 4C, S3E) with the mPTS^PEX22 nest reporter (Figure 4C), supporting the notion that HMGR1 localized to ER nests. Moreover, LDs emerging from these nests retained some HMGR1-positive membrane around them (Figure 4D). Intriguingly, we occasionally observed HMGR1-labeled nests enclosing both LDs and peroxisomes (Figure 4C, 4D).

Expressing non-monomeric fluorescent proteins fused to ER proteins can result in “zippering” of ER membranes^34,35, and HMGR proteins can form similar structures³⁶. However, high signal-capture microscopy of our HMGR1 reporter after pulsed expression showed the protein localizing exclusively to these domains and not in the general ER (Figure S3F, S3G), suggesting that these structures are not artifacts of high levels of self-interacting general ER proteins. Additionally, the punctate HMGR1 reporter localization we observed after pulsed expression matches endogenous HMGR1 localization visualized by immunofluorescence of the native protein in Arabidopsis^36,37. We concluded that HMGR1 specifically sorts to and resides in ER nests.

We also localized squalene epoxidase (SQE1)³⁸ and the oxidosqualene cyclase cycloartenol synthase (CAS1)³⁹, which act in post-squalene sterol synthesis (Figure 4A). Both types of enzymes have been localized to LDs and found in yeast and mammalian LD proteomics⁴⁰, which might reflect ER nest localization. We expressed reporters of SQE1 and CAS1 with markers for peroxisomes and ER nests (Figure S1). In young seedlings, both reporters localized to ER nests enclosing peroxisomes or LDs (Figure 4E–G, S3H, S3I), and after oleate application, both reporters localized to domains around LDs containing the mPTS^PEX22 reporter (Figure S3J, S3K).

To distinguish whether the LD proximity of our nest reporters reflected localization on the LD surface or on surrounding ER nest membrane, we compared SQE1 reporter localization with the localization of OLEOSIN1—a LD coat protein with a long hydrophobic hairpin that extends through the monolayer into the LD core but is incompatible with residence in lipid bilayers⁴¹ that does not concentrate in ER nests (Figure S3L, S3M). Whereas the SQE1 reporter localized primarily to “cups” containing LDs (Figure 4G, 4I), the OLE1 reporter localized evenly around the surface of LDs in young seedlings (Figure 4H, 4I) and after oleate application (Figure S3M). These distinct patterns imply that SQE1 and CAS1 were not directly associated with the LD monolayer but instead localized to residual ER nest membrane that remained with newly-formed LDs (Figure 4J), similar to how LDs forming from HMGR1-labeled nests retained a layer of HMGR-positive membrane (Figure 4D). We concluded that ER nests house sterol biosynthetic enzymes and that organelles born from ER nests can retain some nest membranes partially enclosing them.

RHD3/atlastin functions in ER nest organization

ER nests are complex membrane domains, presumably requiring machinery to organize the membrane and resident enzymes. We examined the influence of atlastin, a dynamin-like GTPase that sculpts ER structure by promoting membrane fusion⁴², on ER nest structure. In animal cells, atlastin also localizes to nascent LDs, and atlastin overexpression enlarges LDs, whereas atlastin deficiency decreases LD size and increases LD numbers⁴³. Atlastin is also implicated in sterol synthesis, as atlastin-deficient neural cells exhibit impaired cholesterol homeostasis⁴⁴.

We expressed ROOT HAIR-DEFECTIVE3 (RHD3), an Arabidopsis atlastin homolog⁴⁵, fused to mTagBFP2 in young seedlings. Like other RHD3 reporters⁴⁶, mTagBFP2-RHD3 localized to the general ER network and connected puncta (Figure S4G). Interestingly, some of these puncta resided within ER nests directly adjacent to sites of budding peroxisomes (Figure 5A). We also localized mTagBFP2 fused to rhd3^T75A, a dominant-negative, GTP-binding-defective variant^46,47. We still observed weak localization of mTagBFP2-rhd3^T75A to nest subdomains (Figure 5B). However, the mutant protein was more dispersed in the general ER, where it caused aberrant ER morphology (Figure S4G), as previously shown⁴⁶.

Figure 5. — (A, B) Induced mTagBFP2-RHD3 (A) and mTagBFP2-rhd3^T75A (B) (blue; arrows) strongly (A) or weakly (B) localize to subdomains of ER nests (green) enclosing peroxisomes (magenta) in young seedlings. Line scans show relative intensities of mTagBFP2-RHD3/rhd3^T75A (blue), mPTS^PEX22-mNG (green), and tdTomato-PTS1 (magenta) in an ER nest (red arrow in merge).

(C) ER nests (arrowheads, labeled by mTagBFP2-HMGR1, blue) have different morphology in 5-day-old wild-type (top) versus *rhd3-7* seedlings (bottom).

(D, E) Oleate-induced LDs (LipidTOX Deep Red; yellow, arrow) are partially enclosed in uniform cup-like ER nests (arrowheads, blue) in wild type (top) whereas ER nests with aberrant and disconnected morphology enclose many small LDs and peroxisomes (green and magenta) (D, bottom) or LDs in *rhd3-7* seedlings (E, bottom).

(F-H) Physiological assays suggest heightened peroxisome function in *rhd3-7*. Bars show median hypocotyl length of seedlings (dots) after 4 days growth in the dark with or without sucrose and IBA (F) or lateral root induction 4 days after transfer of 5-day-old seedlings to media without or with IBA (G). The photo shows seedling roots without or with IBA induction (H). Separate one-way ANOVA tests were performed to compare genotypes on each condition. Letters represent homogenous subsets assigned by Tukey’s posthoc test (P<0.0001).

(I) Peroxisomes are more abundant in the *rhd3-7* mutant. Peroxisome numbers (based on tdT-PTS1 fluorescence) in the elongation zone of 2-day-old seedling roots were quantified from four seedlings of each genotype. Data are shown as mean +/− standard deviations; significance was determined using a one-way t-test.

(J) Following oleate (OA) application, *rhd3-7* displays a reduced triacylglycerol (TAG) to OA ratio. TLC shows wild-type and *rhd3-7* seedling lipids after OA application. Error bars show standard errors of mean ratios of TAG to OA in three biological replicates. Significance was determined using a one-tailed t-test.

See also Figure S4.

We also examined RHD3 localization after inducing LD biogenesis. Without oleate, we observed mTagBFP2-RHD3 puncta in the general ER that often colocalized with mPTS^PEX22-mNG, whereas with oleate, mTagBFP2-RHD3 localized with mPTS^PEX22-mNG-labeled nest membrane around LDs (Figure S4A, Movie S1). mTagBFP2-RHD3 localized in the inner nest membrane layer around oleate-induced LDs, which was presumably not the LD monolayer, as the reporter localized only partially around LDs (Figure S4B, Movie S1), similar to the sterol enzymes (Figure 4G). In contrast, the dominant-negative mTagBFP2-rhd3^T75A reporter localized weakly throughout the general ER of mock-treated seedlings and only partially to the nest membrane around oleate-induced LDs (Figure S4C). Intriguingly, whereas mTagBFP2-RHD3 localized in a continuous sheet around LDs (Figure S4B, Movie S1), mTagBFP2-rhd3^T75A localized to discrete puncta surrounding LDs (Figure S4C, S4D, Movie S2), suggesting discontinuity in the nest membranes remaining around newly-born LDs.

We explored the role of RHD3 by examining ER nest and peroxisome lumen reporters in rhd3-7, an insertional mutant of RHD3 that causes ER structural defects⁴⁵. Pulsed mTagBFP2-HMGR1 expression in older seedlings showed ER nests as small, uniform puncta dispersed throughout cells of wild-type seedlings (Figure 5C). In contrast, nests appeared larger and morphologically aberrant in rhd3-7 (Figure 5C). We observed similar nest morphology using our mPTS^PEX22-mNG and mRuby3-PTS1 reporters to label ER nests and peroxisomes, respectively; ER nests were usually small ER domains that occasionally harbored a peroxisome in wild-type seedlings (Figure S4E) but were misshapen structures containing multiple small peroxisomes in rhd3-7 (Figure S4F). Examining nests using our mTagBFP2-HMGR1 reporter in oleate-treated seedlings showed nests in rhd3-7 roots containing numerous small peroxisomes (Figure S4I) that were not observed in wild-type roots (Figure S4H). We observed similarly disrupted ER nests containing numerous small peroxisomes after oleate application in lines expressing mTagBFP2-rhd3^T75A (Figure S4J), supporting a role for RHD3 in establishing or stabilizing nest structure.

We also observed defects in oleate-induced LD biogenesis from ER nests in rhd3-7. Whereas ER nests were often associated with a single LD in wild-type seedlings, we observed aberrant structures enclosing numerous small LDs in rhd3-7 seedlings that could be aggregated nests (Figure 5D). Additionally, while ER nests around LDs were uniform cup-like structures in wild-type cells, nests around LDs in rhd3-7 were disconnected punctate structures (Figure 5E), similar to the localization of mTagBFP2-rhd3^T75A around LDs (Figure S4D).

To probe how RHD3 deficiency affected peroxisome function, we examined the rhd3-7 mutant in physiological assays. rhd3-7 seedlings grew similarly on media with or without sucrose, a readout of peroxisomal fatty acid β-oxidation, and were sensitive to the peroxisomally-processed auxin precursor, indole-3-butyric acid (IBA), indicating functional peroxisomes (Figure 5F). In fact, rhd3-7 seedlings were more IBA sensitive than wild-type seedlings and formed prolific lateral roots on IBA-supplemented media (Figure 5G, 5H), perhaps because rhd3-7 mutant root cells had more peroxisomes than wild type (Figure 5I).

The ER nest disruption we observed in rhd3-7 might impair the synthesis of LD constituents. We used thin-layer chromatography to examine seedling free fatty acid/TAG profiles after overnight oleate application. Following oleate treatment, rhd3-7 seedlings had similar oleic acid levels but less TAG than wild type (Figure 5J), suggesting reduced TAG biosynthesis or heightened TAG catalysis. We concluded that RHD3/atlastin has a role in organizing ER nest membranes that is important for organelle biogenesis.

ER nests harbor SAR1 isoforms

We also examined SAR1, a small GTPase that orchestrates the assembly and budding of Golgi-bound COPII vesicles from the ER, for a role in ER nests. Cytosolic SAR1 binds target membranes and recruits additional proteins upon activation by its guanine-nucleotide exchange factor⁴⁸. In mammals, Sar1A is primarily involved in canonical ER exit site budding (Figure 3G), whereas Sar1B has additional roles in exporting larger cargo, including lipoproteins⁴⁹. Although some evidence suggests that canonical COPII function does not assist peroxisome formation in mammalian cells⁵⁰, a mammalian COPII protein (Sec16B) is implicated in peroxisome biogenesis⁵¹. Moreover, mammalian Sar1B is found in LD proteomics⁵² and assists in delivering LD coat proteins⁵³. Humans with Sar1B defects have severe cholesterol and TAG homeostasis defects⁵⁴, and while this deficit is ascribed to lipid delivery defects, SAR1-deficient cell lines also show lipid synthesis defects⁵⁴. Arabidopsis has four conserved SAR1 isoforms with functional differences that are not fully elucidated.

We generated plants inducibly expressing fluorescent fusions of SAR1 isoforms and constitutively expressing nest and peroxisome reporters (Figure S1). Remarkably, all four Arabidopsis SAR1 isoform reporters displayed ER nest localization in young seedlings (Figure 6A). We observed many peroxisomes (Figure 6B, Movie S3) and occasional pre-peroxisomes (Figure 6D, S5A) within SAR1-positive ER nests, similar to how we observed LDs enclosed in a layer of nest membrane (Figure 4). While SAR1D appeared to have the most localization to nest membrane around peroxisomes, we also observed the other SAR1 isoforms partially localized around peroxisomes (Figure 6A, S5B).

Interestingly, we observed little to no SAR1-positive nest membrane around peroxisomes packed with membrane (Figure 6B), which we previously found were older peroxisomes that had formed many ILVs³. Because this pattern implies that ER nests might only temporarily enclose peroxisomes after biogenesis, we explored the temporal dynamics of ER nests around peroxisomes. Time-lapse microscopy of peroxisomes enclosed within ER nests over several hours showed SAR1-labeled membranes pulling back from a peroxisome until it cupped the peroxisome before detaching and moving away (Figure 6C, Movie S4). We concluded that SAR1-positive ER nest membranes localized transiently around peroxisomes.

We examined how SAR1 affected ER nests and plant physiology by inducibly expressing dominant-negative sar1^H74L derivatives, which are locked in the active GTP-bound form⁵⁵. Interestingly, these reporters differently affected or localized within ER nests: sar1a^H74L still concentrated in ER nests without notably altering nest morphology; sar1b^H74L had reduced ER nest localization and concentrated in mPTS^PEX22-negative puncta, which may be canonical ER exit sites; and sar1c^H74L and sar1d^H74L both localized to mPTS^PEX22-positive structures that may be disrupted ER nests (Figure S6A). Examining seedlings grown with estradiol to induce sar1 reporter expression showed that sar1a^H74L and sar1b^H74L caused, at most, mild root growth defects, whereas sar1c^H74L and sar1d^H74L arrested seedling development (Figure S6D). However, this exacerbated phenotype may be related to the increased sar1 protein accumulation in these lines (Figure S6B).

We also examined the various SAR1 reporters during LD biogenesis. In mock-treated seedlings, we observed the SAR1B reporter in puncta that did not localize with the mPTS^PEX22 reporter, which may be canonical ER exit sites (Figure S7B). SAR1D had the most substantial nest localization among the isoforms (Figure S6E), and the SAR1D signal in ER nests was brighter than the puncta that likely represent canonical ER exit sites (Figure S7C, S7D). After oleate application, all SAR1 isoforms localized to ER nest membranes around LDs (Figure S6E). The dominant-negative sar1^H74L reporters localized similarly to how they localized during peroxisome biogenesis: sar1a^H74L localized to the nest membrane around LDs without obviously disrupting nest morphology, sar1b^H74L only weakly localized to ER nests, and sar1c^H74L and sar1d^H74L disrupted ER nest morphology (Figure S6F). Interestingly, SAR1 isoforms also partially co-localized with mature peroxisomes (Figure S6E, S6F), hinting at additional peroxisomal roles beyond biogenesis.

To determine if SAR1 expression affected LD biogenesis, we used TLC to monitor conversion of applied oleate to TAG. Seedlings expressing dominant-negative sar1d^H74L had lower TAG to oleic acid ratios than seedlings expressing wild-type SAR1D (Figure S6C), indicating defects in converting free fatty acids to TAG that may reflect LD biogenesis defects. Furthermore, nearly all LDs formed after oleate application were enclosed (fully or partially) by SAR1D-labeled ER nests (Figure 6F). LDs appeared to form from the innermost layer of ER nest membrane (Figure 6E), including nests with smaller LDs surrounded by a layer of SAR1-positive nest membrane and larger LDs partially enclosed in SAR1-labeled nest membrane (Figure 6E, 6F), similar to how SQE1 partially enclosed mature LDs (Figure 4G). We observed similar SAR1-labeled structures in lines not expressing our mPTS^PEX22 nest reporter, with SAR1 reporters localizing in nests around LDs in young seedlings (Figure S5C) and after oleate application (Figure S5A, S5D). Additionally, we observed oleate-induced LDs and pre-peroxisomes within the same ER nests (Figure S5A), supporting the conclusion that these domains are the birthplace of both peroxisomes and LDs. The robust localization of these SAR1 reporters to ER nests, together with the established roles of SAR1 in protein sorting and membrane shaping in other systems, suggests that SAR1 isoforms may function in nest protein sorting and/or membrane shaping.

ER nests remaining around peroxisomes appear to control ILV formation

The residual nest membrane layer around peroxisomes and LDs emerging from nests might provide a means to regulate contact between these organelles. Intriguingly, we often observed ILVs restricted to peroxisomal regions coinciding with regions where nest membrane was absent (Figure 7A, 7B), suggesting that remnant nest membrane might regulate its progeny organelles. Peroxisomal ILV formation is associated with the trafficking of fatty acids from LDs to peroxisomes, when ILVs appear to form directly at peroxisome-LD contact sites³. We observed accumulated ILVs at the regions of peroxisomes not enclosed by ER nests that were contacting LDs (Figure 7C), suggesting that ER nests might define contact sites between peroxisomes and LDs. We explored this relationship using TEM of wild-type seedlings (lacking reporters) following oleate-induced LD formation. In agreement with our reporter data, we observed smooth ER membranes (presumed ER nests) around LDs and peroxisomes that were excluded from contact sites between peroxisomes and LDs (Figure 7D). Furthermore, we observed peroxisomal ILVs forming directly at these LD contact sites defined by the surrounding nest membrane (Figure 7E).

Figure 7. — (A, B) Examples of SAR1D-mTagBFP2 (blue) enclosing mature peroxisomes (green and magenta) in which the SAR1D reporter is absent from sites of ILV formation (arrows).

(C) An example of SAR1D-mTagBFP2 (blue) enclosing a mature peroxisome (green and magenta) in contact with a LD (LipidTOX Deep Red; yellow). Box shows a magnification of the peroxisomal ILVs (green) forming at the LD contact site, which lacks SAR1D-labeled nest.

(D, E) TEM images of an oleate-treated wild-type seedling (without reporters) showing LD-peroxisome (P) contact sites (arrows) where apparent nest membranes (arrowheads) are absent (D) and a peroxisome forming an ILV at a LD contact site (arrow) flanked by apparent nest membrane (arrowheads) (E).

(F) Stills from time-lapse Movie S9 of hypocotyl cells (which catabolize fats in the dark to fuel elongation) showing ER nests (SAR1D-mTagBFP2, blue) moving to and enclosing peroxisomes (magenta). The peroxisomes (arrow) enclosed by ER nests enlarge as the ER nests (arrowhead) shrink, suggesting lipid transfer to peroxisomes.

(G) Image slices of the indicated channels from three timepoints of panel F show the ER nest (arrowhead, blue) contacting and engulfing peroxisomes (green and magenta, white arrow), which contact a LD (brightfield, yellow arrow) as the peroxisomes enlarge.

(H) Working model for ER nest dynamics. Peroxisomes (and lipid droplets) form from the inner layer of ER nests, which can remain partially enclosing the nascent organelles or eventually retract. When metabolic demand increases, ER nests may return to support peroxisomes (and lipid droplets).

See also Figure S8.

ER nests are dynamic and can re-enclose organelles during heightened metabolism

To investigate the temporal dynamics of ER nests, we used time-lapse confocal microscopy to image seedlings expressing the mTagBFP2-SAR1D reporter. This analysis showed instances of larger ER nests of young seedlings shrinking (Figure S8A, Movie S5), ER nests fusing (Figure S8B, Movie S6), ER nests dividing (Figure S8C, Movie S7), and ER nest morphology changing from a punctate structure to a flat sheet (Figure S8D, Movie S8). These events appeared in ER nests that were not enclosing or in contact with visible peroxisomes or LDs and suggest that ER nests are dynamic structures with some features of independent organelles.

Imaging 3-day-old light-grown seedling hypocotyls showed that most peroxisomes were not enclosed in ER nests at the start of imaging. However, as the imaging continued overnight, we observed ER nests moving to and enclosing peroxisomes as the peroxisomes expanded (Figure 7F, 7G, Movie S9. Hypocotyl LDs fuel cell expansion in the dark but appear to be catabolized more slowly than cotyledon LDs in light-grown seedlings. We speculate that hypocotyl LD mobilization increased in the darkness of the imaging room to fuel cell elongation as the seedlings sought to escape the darkness. This movement of ER nests to peroxisomes in seedlings shifted to darkness hints that nests support these organelles as they participate in heightened lipid mobilization (Figure 7H).

DISCUSSION

We have characterized the ER domains (ER nests) where peroxisomes and LDs are born. These complex domains exclude general ER membrane proteins, accumulate lipid-synthesizing enzymes, and are shaped by RHD3/atlastin and SAR1. ER nests appear to function as non-canonical ER exit sites, where instead of budding Golgi-bound vesicles with lumenal cargo (Figure 3G), distinct organelles are born (Figure 3H, 4J). These domains can provide contact sites with progeny organelles, restrict contact with other organelles, and move back to and enclose organelles. Beyond organelle biogenesis, ER nests are persistent and dynamic ER subdomains where lipid synthesis may be sub-compartmentalized. We observed dramatic changes in ER nest size (Figure 6C, S8A) and morphology, including nests dividing, fusing, and transitioning from punctate to sheet-like structure (Figure S8). Similar ER structures have been observed enclosing nascent LDs in human cells transitioning to adipose cells⁵⁶, enclosing nascent peroxisomes in mammalian^57,58 and yeast cells⁵⁹, and in animal cells with high sterol synthesis⁶⁰, suggesting that ER nests are present throughout eukaryotes.

ER nests exhibit robust protein sorting. The accumulation of PEX22-based reporters, sterol enzymes, and SAR1 (Figure 1, 4, 6, S3) but near total exclusion of a general ER membrane reporter (Figure 1E, 1F) indicate segregation of nest proteins from general ER proteins. The nest and ER reporters we utilized—mPTS^PEX22 and the TMD of CB5D—included only the transmembrane and juxtamembrane domains of their respective proteins. Perhaps a difference in lipid composition contributes to partitioning these transmembrane domains into nests. For example, sterol-rich membranes accumulate specific proteins while excluding others⁶¹, and sterols are important for mammalian peroxisome formation⁶² and targeting coat proteins to plant LDs⁶³. The sterol biosynthetic enzymes we observed concentrating in ER nests (Figure 4) might hint that local sterol synthesis could impact nest protein sorting. Apart from the PEX22 reporter, we did not observe other nest reporters (e.g., HMGR1) localizing directly on peroxisome membranes (Figure 4B, 4D), suggesting a sorting mechanism within ER nests to ensure budding peroxisomes do not acquire nest proteins.

Peroxisome and LD biogenesis presumably requires stringent sorting to avoid contaminating the ER. Peroxisomal enzymes often produce oxidative byproducts, which could be toxic in the redox-sensitive environment of the ER lumen. Similarly, TAG biosynthetic enzymes produce lipotoxic intermediates (e.g., free fatty acids) that might destabilize general ER membranes. Spatially restricting lipid synthesis would protect the general ER from this lipotoxicity while increasing metabolic efficiency. The multilamellar structure of ER nests may also enhance lipid biosynthesis, as generating multilamellar membranes in engineered microbes increases squalene production⁶⁴.

We occasionally observed peroxisomes and LDs within the same ER nests (Figure 2E, 4D, S5A), supporting previous demonstrations of overlapping biogenesis machinery of these two organelles. For example, yeast Pex30, initially characterized for its role in regulating peroxisome numbers⁶⁵, also participates in LD biogenesis¹³. Moreover, seipin pex30 double mutants accumulate ER membrane whorls instead of LDs upon oleate application, and these whorls, like the ER nests described here, exclude a general ER reporter⁶⁶. Interestingly, peroxisome biogenesis is compromised in seipin pex30 cells⁶⁶, hinting that these whorls may be dysfunctional ER nests and that defects in the biogenesis of one nest-derived organelle might impact the formation of others.

We speculate that peroxisome biogenesis involves peroxisomal proteins sorting to the inner layer of ER nests, allowing lumenal protein import from the cytosol into the growing pre-peroxisomal bud to drive nascent peroxisome expansion (Figure 3H). This model requires continuity of the inner nest membrane with the cytosol, where lumenal proteins are synthesized. Mammalian cells with import defects have fewer peroxisomes visible with a peroxisomal membrane protein antibody⁶⁷, and yeast mutants lacking PEX13, a critical component of the peroxisomal import machinery, display peroxisomes stuck in multilamellar structures⁶⁸. This peroxisome biogenesis model is analogous to our model for LD biogenesis in ER nests (Figure 2H), except that protein import rather than lipid synthesis drives organelle growth. Intriguingly, a yeast lipin mutant, defective in a phospholipase that converts phospholipids into diacylglycerol for TAG formation, displays small LDs enclosed in thick ER⁶⁹. Perhaps lipin mobilizes excess nest membrane into TAG as LDs mature.

We detected a small amount of the peroxisomal mPTS^PEX26 reporter in ER nests after several days of expression (Figure 3E), suggesting that membrane proteins that escape the budding process might remain marooned in the nest. Unlike in wild type, in a pex5 mutant, we observed abundant nest localization of the mPTS^PEX26 reporter (Figure 3C, 3D) and even in the general ER (Figure 3F), suggesting that peroxisomal membrane proteins can mislocalize to the general ER when lumenal protein import is impeded. Mislocalization of peroxins to mitochondria contributes to human disorders caused by peroxisome dysfunction⁷⁰, and perhaps mistargeting of peroxisomal proteins to the ER also contributes to these disorders.

The only known functions of PEX22 are to anchor and activate the ubiquitin-conjugating enzyme PEX4 for ubiquitinating peroxisomal proteins⁷¹, but our PEX22 reporter localization suggests that PEX4 might have additional substrates in ER nests and perhaps in the general ER. For example, the size and protein composition of ER subdomains might be adjusted by ubiquitin-directed proteolysis of resident membrane proteins. Self-interacting ER membrane proteins can induce membrane whorls, which have a morphology similar to ER nests, and various sterol biosynthetic enzymes interact with one another⁷². Thus, nest proteins may recruit or stabilize additional nest membrane layers, and degrading these proteins would allow nests to shrink, thereby tuning capacity for lipid synthesis (which involves reactions occurring in the lipid bilayer). Indeed, many LD-related proteins are regulated by ubiquitination and proteasomal degradation⁷³, and whorls of smooth ER (often enclosing LDs) are an identifying feature of cells with high lipid-synthesizing loads, such as sterol-producing Leydig cells^60,74. We observed that the large ER nests in young seedlings (where peroxisome biogenesis is high) shrank as seedlings aged and peroxisome biogenesis decreased (Figure S8A, Movie S5). Ubiquitination might also trigger removal of errant nest proteins from the general ER. Similarly, ubiquitin-dependent degradation of enzymes in the general ER enforces the LD localization of some lipid-synthesizing enzymes that sort from the ER to LDs⁷⁵. In contrast to our PEX22 reporter, which localized to the general ER after pulsed expression (Figure 1D, S2B), several other fluorescently labeled enzymes were specific to ER nests (Figure S3F–K). Thus, ubiquitination via PEX4 and PEX22 might control nest proteins in and out of ER nests.

Reporters for TAG and sterol biosynthetic enzymes, including HMGR, localized in ER nests (Figure 2D, 4). Despite being considered an ER protein, HMGR localizes in mysterious puncta in plants³⁷ and mammalian cells⁷⁶. Although these structures have been postulated to be aberrant structures caused by the “zippering” effect of highly expressed oligomerizing ER membrane proteins, including non-monomeric fluorescent protein reporters³⁴, endogenous HMGR1 also concentrates in ER puncta³⁷, and we did not detect HMGR in the general ER even after pulsed expression (Figure S3D–G), suggesting that these puncta report normal HMGR localization. HMGR ubiquitination controls sterol production⁷⁷, and ubiquitination of HMGR in the general ER could restrict HMGR activity to ER nests.

Reporters for the sterol biosynthetic enzymes SQE1 and CAS1 also localized in ER nests, predominantly in nest membranes that partially surrounded LDs (Figure 4E–G). Although related enzymes in other systems appear to sort to LDs via the ER⁴⁰, whether these enzymes are localized to the LD monolayer or ER membrane abutting LDs remains controversial. Photo-bleaching experiments indicate that some lipid-synthesizing enzymes are in membranes continuous with the ER⁷⁸, and LDs visualized via freeze-fracture EM appear to be enclosed in a sheet of ER and can have cup-like ER partially around them⁷⁹. The large size of Arabidopsis organelles affords the resolution to demonstrate that these enzymes usually do not entirely surround a LD, but instead localize to mPTS^PEX22-positive membrane around LDs that is distinct from the localization of the LD coat protein OLE1 (Figure 4G–I). Moreover, we observed LDs associated with a cup of SAR1-labelled nest membrane (Figure 6E, 6F) and LDs enclosed in ER using an ER-lumen reporter (Figure S2G–I), indicating ER around LDs rather than on the LD monolayer.

Our data provide mechanistic insights into the nest assembly and budding process. Biogenesis defects of both peroxisomes and LDs were apparent upon RHD3/atlastin disruption (Figure 5, S4). Perhaps RHD3 acting as an ER fusogen⁴² indirectly maintains ER nest morphology. However, RHD3 localization in nest subdomains close to budding peroxisomes or LDs (Figure 5) suggests a direct role for RHD3 in nest function. Atlastin-mediated fusion may help establish ER nests or nest subdomains. Moreover, atlastin participates in COPII assembly independently of its fusion activity⁸⁰. Because we implicated the COPII component SAR1 with ER nests (Figure 6), a non-fusion function of RHD3 may additionally or alternatively be important for ER nests. Disrupting atlastin in animal cells results in numerous fragmented LDs⁴³. RHD3 may help organize the innermost layer of ER nests where nascent organelles emerge. For example, we readily observed ER nests fusing (Figure S8B, Movie S6), and RHD3 may promote the continuity of fusing ER nest membranes. Thus, the dispersed budding machinery of disorganized nests may produce more and/or smaller peroxisomes and LDs in the rhd3-7 null mutant (Figure 5).

Finding the canonical ER-budding component, SAR1, in ER nests (Figure 6) might implicate COPII function with nests. Dominant-negative SAR1 variants altered ER nest structure (Figure S6A) and arrested seedling development (Figure S6D), suggesting that SAR1 shapes ER nest membranes and hinting that ER nests may be necessary for plant growth and development. The role of mammalian Sar1B in targeting some LD proteins⁵³, its localization to liver peroxisomes⁸¹, and its connection to lipid synthesis⁵⁴ support an ER nest function for SAR1. Additionally, the interactome of human SAR1B includes several lipid-synthesizing enzymes, including HMGR and the peroxisomal protein PEX11B⁸², underscoring the connection between ER nests, lipid synthesis, and peroxisomes. The SAR1-labeled membranes we observed around peroxisomes, pre-peroxisomes (Figure 6B, 6D), and LDs (Figure 6E, 6F) suggest that both organelles form from the inner layer of ER nests.

ER nests appeared to regulate the organelles that they enclosed, adding to the ways in which the ER controls the dynamics of other organelles^11,12. We observed organelles either nestled in a cup of nest membrane or mostly enclosed by the membrane (Figure 6). We observed this residual nest membrane retracting from organelles and reforming into a circular structure that moves away (Figure 6C, Movie S4). It will be interesting to learn whether these structures remain functional as nest membranes that could birth additional organelles. Intriguingly, we observed ER nest membrane defining sites of peroxisomal ILV formation and restricting peroxisome-LD contact sites (Figure 7A–E), suggesting that ER nests spatially regulate peroxisome and LD function. Peroxisomal metabolism usually involves the transfer of intermediates to and from other organelles, with peroxisomes directly contacting partner organelles⁸³, and it will be interesting to explore how ER nests might influence these interactions. For instance, regulating contact sites may be important for lipid synthesis. As LD fats are mobilized to peroxisomes, having ER nest surrounding the contact site could prevent any released fatty acids that escape peroxisomal import from diffusing into the rest of the cytoplasm (where they could exert lipotoxicity). Indeed, TAG-synthesizing enzymes protect adipocytes undergoing lipolysis from ER stress⁸⁴. Although TAG synthesis during conditions of TAG catabolism seems paradoxical, perhaps escaped fatty acids are captured by ER nests (which house TAG synthesizing enzymes) and converted back into TAG to ensure that fatty acids are transferred only at contact sites. Beyond involvement in lipid metabolism, it will be interesting to learn whether ER nests envelop and regulate organelles in addition to peroxisomes and LDs.

Additional structures might hail from ER nests. Intriguingly, autophagosome biogenesis relies on components implicated in ER nest function. For example, Arabidopsis autophagosome biogenesis requires SAR1D⁸⁵, and nascent autophagosomes are enveloped in ATG5-labeled ER structures⁸⁶. Furthermore, mammalian atlastin organizes autophagosome biogenesis⁸⁷, and Sar1 disruption impairs autophagosome biogenesis⁸⁸. In animal cells, autophagosome biogenesis appears to involve the ER⁸⁹, and autophagosomes originate within discrete ER-connected structures that exclude general ER proteins (based on a cytochrome B reporter)⁹⁰. Intriguingly, the protein DFCP1 has a prominent role in this ER subdomain⁹⁰ and is also important for LD biogenesis and localizes around nascent LDs⁹¹, which might reflect a role within ER nests. Additionally, DFCP1 regulates the ATGL-mediated lipolysis of LDs, which could connect to how we observe ER nests mediating peroxisome-LD contacts, particularly because ATGL is brought to LDs from peroxisomes⁹². It will be interesting to learn whether autophagosomes or other structures arise from ER nests. Finally, ER nests may be viral budding platforms. Several viruses replicate using LDs⁹³, and viral proteins localize around nascent LDs or ER membranes around LDs. For instance, SARS-CoV-2 replicates in mysterious multilamellar ER structures containing atlastin, but not general ER proteins, that associate with LDs⁹⁴. Additionally, several viruses, including hepatitis C and SARS-CoV-2, downregulate peroxisome biogenesis to evade peroxisome-mediated immune signaling⁹⁵, and SARS-CoV-2 downregulates autophagosome biogenesis⁹⁶, which might be mediated at ER nests.

Arabidopsis is a powerful system to investigate ER nests and their progeny organelles. The large size of Arabidopsis cells and organelles allows sub-organellar visualization of these structures in living, intact plants. In other systems, these might be mistaken for other cellular components. For instance, the exclusion of general ER membrane proteins from nests would make them resemble other organelles. Multilamellar membranes enclosing organelles could appear similar to lysosomes or autophagosomes under electron microscopy. The role of ER membranes in restricting peroxisomal ILVs has perhaps gone undiscovered because peroxisomes in other models are usually too small to observe intraorganellar detail using light microscopy and can be difficult to identify with electron microscopy due to poor membrane resolution and structural similarity to lysosomes or multivesicular endosomes. Finally, the dynamism of ER nests may help explain why they have eluded identification until now.

Limitations of the study

We could not visualize the earliest steps of peroxisome or LD biogenesis due to the resolution limitations of light microscopy and the difficulty of peroxisome identification using TEM. Lipid droplets were induced with oleate, which is not a condition experienced by plants in the wild. Expression of reporters could impact the morphology or function of the targeted organelles. Finally, this study was limited to Arabidopsis seedlings. Because efficient LD and peroxisome biogenesis correlate with increased lifespan and fitness in C. elegans¹⁴, and dysregulated lipid metabolism is observed in various cancers⁹⁷, it will be interesting to learn how these findings apply in other eukaryotes.

RESOURCE AVAILABILITY

Lead contact

Requests for further information and resources should be directed to and will be fulfilled by the lead contact, Bonnie Bartel (bartel@rice.edu).

Materials availability

Requests for unique biological materials (e.g., plant lines, DNA constructs) should be directed to and will be fulfilled by the lead contact.

Data and code availability

The data supporting the findings are in the main text or supplementary information.
This paper does not report original code.
Any additional information required to reanalyze the data reported in this paper is available from the lead contact upon request.

STAR METHODS

EXPERIMENTAL MODEL AND STUDY PARTICIPANT DETAILS

Arabidopsis

Transgenic lines and mutants were in the Arabidopsis thaliana Colombia-0 (Col-0) accession. Plants were grown under continuous white light at 22°C unless otherwise indicated. Seeds were surface-sterilized and stratified at 4°C for 1–3 days before being sown on plant nutrient (PN) medium⁹⁸ supplemented with 1% sucrose (PNS) and solidified with 0.6% or 1% agar for horizontal or vertical growth, respectively. Six- to 12-day-old seedlings were transplanted to soil for seed production.

METHOD DETAILS

DNA cloning methods

Constructs were derived from a modular tri-fluorescent construct synthesized by GenScript, containing three different sets of a promoter driving a fluorescent protein with a distinct terminator, and all pieces flanked by a unique pair of restriction sites to create either pUC57–35S-mTagBFP2-HA, ACT2-TdTomato-PTS1, UBQ10-mNG-mPTS^PEX26 or pUC57–35S-mTagBFP2-HA, ACT2-TdTomato-PTS1, UBQ10-mPTS^PEX22-mNG. mPTS^PEX22 and mPTS^PEX26 sequences were chosen from constructs generated previously³, with mPTS^PEX22 including the first 94 amino acids of PEX22 and mPTS^PEX26 including the last 90 amino acids of PEX26. Construct variants were generated using restriction sites to drop out pieces and Gibson Assembly⁹⁹ to clone in a PCR-generated insert. Other cDNA sequences used were obtained through PCR of reverse-transcribed Arabidopsis seedling RNA. The CB5DTMD reporter included the last 40 amino acids of CB5D, and all other constructs included the entire cDNA open reading frame. Prior to plant transformation, reporter plasmid inserts were recombined into pMCS-GW¹⁰⁰ (for reporters with all promoters encoded in the insert fragment) or into pFZ19¹⁰¹ (for estradiol-inducible expression of the mTagBFP2-containing reporter in the 5’-most position in the transgene) via Gateway recombination¹⁰² using LR clonase II recombinase (Invitrogen) after excising the 35S promoter in the 5’ position of the mTagBFP2-containing reporter via XhoI digest. Plasmid 3516 (Figure S1) was generated by excising the UBQ10-mRuby3-PTS1 sequence of plasmid 3408 with SpeI and NheI. Plasmids were verified by sequencing via assembled Oxford Nanopore reads by Plasmidsaurus (Eugene, OR).

Arabidopsis transformation

DNA constructs were transformed into Agrobacterium tumefaciens strain GV3101 (bearing the PMP90 helper plasmid¹⁰³ using electroporation, and the resultant transformed Agrobacterium strains were used to transform wild-type Arabidopsis plants using the floral dip method¹⁰⁴. Depending on the construct, transformants were selected using herbicide resistance and examination of fluorescence using a Leica MZ16FA fluorescent stereomicroscope. Plants transformed with constructs conferring Basta resistance were selected following growth on PNS plates containing 10 μg/mL Basta for several days with plants fluorescent for all three constructs being selected from the Basta-resistant plants in the T₁ generation. Plants transformed with constructs not bearing a Basta resistance gene (lines containing estradiol-inducible constructs) were selected by fluorescence alone following growth on PNS containing 25 μM β-estradiol for 2 or 3 days. All selection media also contained 20 μg/mL timentin to prevent Agrobacterium growth. Selected plants were moved to PNS until large enough to move to soil for seed production.

Estradiol induction

To express genes under the control of the estradiol-inducible promoter¹⁰⁵, plants were either sown on solid PNS media supplemented with 25 μM β-estradiol (from a 100 mM stock dissolved in DMSO) and grown on vertically-positioned plates before imaging, or seedlings were grown on solid PNS media without estradiol for a set number of days and then transferred to liquid 1/6x PNS supplemented with 200 μM Tween-20 and either DMSO (mock) or 25 μM β-estradiol.

Lipid droplet induction

To induce LD formation with exogenous oleate, seedlings were grown on solid PNS media for a set number of days before being transferred to liquid 1/6x PNS medium supplemented with 200 μM Tween-20 and either DMSO (mock) or 0.05% oleic acid (from a 10% stock dissolved in DMSO).

Immunoblotting

Seedlings were grown vertically in the light on PNS for 4 d and then treated for 1 day in liquid medium supplemented with 25 μM estradiol and mock or 0.05% oleic acid. Protein was extracted by grinding frozen plant tissue, adding 3 volumes of sample buffer [500 mM Tris pH 8.0, 4% (w/v) lithium dodecyl sulfate, 1 mM EDTA, 20% (w/v) glycerol, 11 μM Coomassie blue G250, 16.6 μM phenol red, 50 μM dithiothreitol (DTT)], boiling at 100 °C for 5 minutes, and centrifuging to remove cell debris. Protein extracts (8 μL) were loaded on Bolt 10% (w/v) Bis-Tris gels (Invitrogen) alongside equal volumes of prestained protein markers (NEB P7719S), electrophoresed in 50 mM MOPS running buffer (50 mM MOPS free acid, 50 mM Tris base, 0.1% (w/v) SDS, 1 mM EDTA), and transferred to Hybond-ECL nitrocellulose membranes (Amersham, Protran Premium 0.45 μm NC 10600003) with an eBlot L1 transfer system (GenScript). After transfer, membranes were air-dried at room temperature for one hour followed by overnight blocking in 8% Carnation instant non-fat dry milk in TBST (20 mM Tris pH 7.5, 150 mM NaCl, 0.1% (v/v) Tween-20) at 4°C with rocking. After blocking, membranes were incubated with primary antibodies diluted in 8% milk in TBST overnight at 4°C with rocking. Primary antibodies were mouse anti-HSC70 (1:50,000; Stressgen SPA-817), rat anti-HA (1:100; Roche clone 3F10), rabbit anti-PEX14 (1:10,000; Agrisera AS08 372), and rabbit anti-PMDH2¹⁰⁶ (1:5000) and were incubated overnight at 4°C. After incubation with primary antibodies, membranes were washed with three 5-minute washes in TBST, and then incubated with horseradish peroxidase (HRP)-linked goat anti-mouse (1:5000), goat anti-rat (1:2500), or goat anti-rabbit (1:5000) secondary antibodies for 2–4 hours at 4°C with rocking. Membranes were imaged with WesternSure Premium Chemiluminescent substrate (Fisher, 50–489–552) using an Odyssey Fc imaging system (LI-COR, 2801–02). Membranes were serially incubated with different primary antibodies without stripping.

Confocal microscopy

Seedlings were imaged using live-cell confocal fluorescence confocal microscopy after being grown in the light on solid PNS media. Plants with estradiol-inducible reporters were grown on PNS media supplemented with 25 μM β-estradiol (for imaging 2- or 3-day-old seedlings) or were grown on media without estradiol and transferred to liquid PNS media with 25 μM β-estradiol and incubated overnight before imaging. For regular confocal imaging, seedlings were mounted in water or LD dye solution on glass slides (VWR; 48311–950) with 0.16-mm coverslips (VWR; 48393–241). For time-lapse confocal imaging, a thin layer of agar was set into an imaging well slide by placing 1 mL of warm 0.1% agar PNS media into the well slide and allowing it to solidify, and then excised cotyledons or hypocotyls were placed underneath the slab for imaging. For most experiments, fluorescence was captured using an Olympus FV3000RS inverted laser scanning confocal microscope equipped with Fluoview Acquisition software (version 2.4.1.198), a UPLXAPO 60x/1.42 oil-immersion objective, and standard multialkali spectral GaAsP detectors.

To stain LDs in seedlings without a blue-fluorescent reporter, seedlings were incubated in 100 μM monodansylpentane (MDH) in 50 mM Tris (pH 8) for at least 20 minutes. To stain LDs in seedlings expressing a blue-fluorescent reporter, seedlings were incubated with LipidTOX Deep Red diluted 1:200 in 50 mM Tris (pH 8) for at least 30 minutes. MDH fluorescence was excited with a 401-nm laser and captured at 645–700 nm. LipidTOX Deep Red fluorescence was excited with a 640-nm laser and captured at 645–700 nm. Due to overlap with chloroplast autofluorescence, LipidTOX Deep Red signal that was close to abundant chloroplasts was not used and instead LDs were shown in the brightfield channel based on the difference in diffraction of light.

To visualize the various fluorescent reporters, mTagBFP2 fluorescence was excited with a 405-nm laser and captured at 430–470 nm, mNeonGreen fluorescence was excited with a 488-nm laser and captured at 500–540 nm, and tdTomato fluorescence was excited with a 561-nm laser and captured at 570–600 nm. All three channels were imaged on separate imaging tracks, which switched by pixel line. When LipidTOX Deep Red dye was also imaged, it was imaged on the same track as mTagBFP2.

Images were acquired as z-stacks, and displayed images are slices selected from z-stacks unless specified as a z-projection. All images (except Figure S2F) were acquired with 16-bit depth at 512×512 resolution and no averaging. Figure S2F was acquired with 12-bit imaging at 1024×1024 resolution and was an average of two scans.

Endosomes (Figure S2F) were visualized by incubating seedlings expressing mPTS^PEX22-mNG with 5 μM FM4–64 dye in darkness for 90 minutes. Seedlings were imaged using a Carl Zeiss 710 confocal microscope equipped with Zen 2010 software (version 6.0.0.485), a Meta detector ×100/1.4 oil-immersion objectives. FM4–64 fluorescence was excited with a 543-nm laser and captured at 592–758 nm, and mNeonGreen was excited with a 488-nm laser and captured at 590–561 nm.

Transmission electron microscopy

Freshly collected samples were fixed overnight in Karnovsky’s fixative, secondarily fixed in 1% osmium tetroxide for one hour, dehydrated in a graded series of ethanol, and embedded in epoxy resin. Samples were sectioned at 100 nm thickness using a Leica EM UC7 ultramicrotome and then stained with uranyl acetate and Reynold’s lead citrate. Sections were examined using a JEOL JEM-1400Flash TEM equipped with a high contrast pole piece and an AMT NanoSprint15 Mk-II sCMOS camera.

Lipid analysis

To monitor TAG formation from applied oleic acid, seedling lipids were extracted using the Folch extraction method^63,107 and separated by thin-layer chromatography (TLC). Approximately 25 seeds were grown in liquid 1/6X PNS for 5 days before LD induction with oleate for 1 day. Seedlings were frozen in liquid nitrogen and ground in 1.7-mL centrifuge tubes with a plastic pestle. 500 μL of chloroform/methanol/formic acid (1/2/0.1, by volume) were added and samples were vortexed for at least 10 minutes at room temperature to extract lipids. 0.5 volumes of phase separation solution (1M KCl and 0.2 M H₃PO₄) were added, samples were vortexed, and samples were centrifuged at 13,000 rpm for at 5 minutes. Equal volumes of the lower phase were spotted onto TLC plates (silica gel 60, Sigma, 1.05721) and separated by thin-layer chromatography using a solvent system consisting of hexane/diethyl ether/acetic acid (70/30/1, by volume). TAG was visualized by dipping TLC plates into 5% H₂SO₄, drying at room temperature, and heating at approximately 100 °C for 5–10 minutes until color developed. Control lipids (5–10 μL) containing a mixture of 0.5% TAG (glyceryl trioleate, Sigma 92860), 0.2% DAG (dioleoylglycerol, Sigma D8894), and 0.1% oleic acid (Sigma O1008) dissolved in chloroform were spotted on each TLC plate to allow assignment of TAG and oleic acid in the experimental samples.

QUANTIFICATION AND STATISTICAL ANALYSIS

All fluorescent reporters were visualized in multiple seedlings in the T₁ generation (primary transformants) to survey multiple independent insertion events and ensure that observed events were not dependent on particular insertion sites in the genome. Selected confocal images for each construct represent events that were observed in multiple cells from at least three different seedlings in the T₂ or T₃ generation.

Peroxisome and LD size and number were quantified by taking z-projections of LD or peroxisome channels made in FIJI (version 1.54f) from three or four seedlings and importing them into Ilastik¹⁰⁸ (version 1.3.2) to create binary masks by training the pixel classification method to the respective channel without or with oleate treatment. The masks were then imported back into FIJI, and the watershed function and close function were used to separate touching objects and remove single-pixel noise, respectively. Peroxisome cross-sectional area was then measured in FIJI and converted to diameter in Microsoft Excel. Data were graphed in PRISM (version 10.0.3).

TLC plates were scanned using a flatbed scanner. TIF images were imported into Image Studio software (version 5.2.5) for quantification of TAG and oleate signals. Data were graphed using PRISM (version 10.0.3).

Arabidopsis root and hypocotyl lengths were quantified by growing plants vertically before scanning plates with a flatbed scanner. Scans were imported into FIJI, the scale was set to match an included ruler in the image, and root or hypocotyl length was measured manually in FIJI. To quantify lateral roots per root length (Figure 5G), lateral root number and root length were counted using a Leica MZ16FA stereomicroscope to accurately quantify the numerous, small lateral roots of rhd3-7. Data were graphed using PRISM (version 10.0.3).

Statistics were performed using JMP Pro (version 17.0.0). One-way t-tests were used to compare organelle numbers (Figure 3, 5I), and one-tailed t-tests were used to compare ratios of triacylglycerol to oleate (Figure 5J, S6C). One-way ANOVA was performed to compare IBA responsiveness of rhd3-7 (Figure 5F, 5G) and root lengths of SAR1 reporter-expressing seedlings (Fig. S6D). When P-values were <0.05, Tukey’s posthoc test was used to compare means and assign homogenous subsets. Figures were assembled in Adobe Illustrator (version 28.0).

Supplementary Material

Movie S1. RHD3 localizes in subdomains of ER nests around oleate-induced lipid droplets, related to Figure 5 and S4A–B.

mTagBFP2-RHD3 (blue) localizes to sheets partially enclosing oleate-induced LDs (brightfield) within ER nests (mPTS^PEX22-mNG; green) in hypocotyl cells of 6-day-old seedlings incubated with oleate overnight. Peroxisomes (tdT-PTS1; magenta) are also shown. Video shows images from hypocotyl epidermal cells extending into the tissue at 1 μm intervals. Scale bar is 10 μm.

Download video file^{(5.5MB, mp4)}

Movie S2. rhd3^T75A localizes in puncta around lipid droplets, related to Figure 5 and S4C–D.

mTagBFP2-rhd3^T75A (blue) localizes to puncta around oleate-induced LDs (brightfield) within ER nests (mPTS^PEX22-mNG; green) in hypocotyl cells of 6-day-old seedlings incubated with oleate overnight. Peroxisomes (tdT-PTS1; magenta) are also shown. Video shows images from the outside of cotyledon epidermal cells extending to the beginning of underlying mesophyll cells at 1 μm intervals. Scale bar is 10 μm.

Download video file^{(3.7MB, mp4)}

Movie S3. SAR1D localizes to ER nest membrane around young peroxisomes, related to Figure 6B.

SAR1D-mTagBFP2 (blue) localizes to ER nest membrane partially enclosing peroxisomes (tdT-PTS1; magenta) in cotyledon cells of 3-day-old seedlings. Video shows images from near the top to near the bottom of cotyledon epidermal cells at 1 μm intervals. Scale bar is 10 μm.

Download video file^{(2.2MB, mp4)}

Movie S4. ER nests can leave peroxisomes, related to Figure 6C.

ER nest membrane around peroxisomes (tdT-PTS1; magenta) that is positive for SAR1D-mTagBFP2 (blue) initially surrounds peroxisomes but retracts over time before fully moving away in cotyledon cells of 4-day-old seedlings. Video shows z-projections (1 μm stack intervals through cotyledon epidermal cells) acquired every 15 minutes over 6 hours. Scale bar is 10 μm.

Download video file^{(2MB, mp4)}

Movie S5. ER nests can shrink after periods of high activity are concluded, related to Figure 6 and Figure S8A.

Two ER nests labelled with SAR1D-mTagBFP2 (blue) imaged over time in cotyledon cells of 4-day-old seedlings shrinking in size over several hours. Video shows z-projections (1 μm stack intervals through cotyledon epidermal cells) acquired every 15 minutes over 15 hours. Scale bar is 10 μm.

Download video file^{(3.5MB, mp4)}

Movie S6. ER nests can fuse together, related to Figure 7 and S8B.

Two ER nests labelled with SAR1D-mTagBFP2 (blue) imaged over time in hypocotyl cells of 3-day-old seedlings fusing together. Video shows z-projections (1 μm stack intervals through hypocotyl epidermal cells) acquired every 15 minutes over 4 hours. Scale bar is 5 μm.

Download video file^{(1.3MB, mp4)}

Movie S7. ER nests can divide, related to Figure 7 and S8C.

ER nests labelled with SAR1D-mTagBFP2 (blue) imaged over time in hypocotyl cells of 3-day-old seedlings shows an extension coming off an ER nests that eventually separates and moves away. Video shows z-projections (1 μm stack intervals through hypocotyl epidermal cells) acquired every 15 minutes over 2.5 hours. Scale bar is 5 μm.

Download video file^{(810.4KB, mp4)}

Movie S8. ER nests can change their morphology and flatten into sheets, related to Figure 7 and S8D.

ER nests labelled with SAR1D-mTagBFP2 (blue) imaged over time in hypocotyl cells of 3-day-old seedlings shows an ER nest with a punctate morphology flattening into a sheet of membrane that then moves within the cell. Video shows z-projections (1 μm stack intervals through hypocotyl epidermal cells) acquired every 10 minutes over 20 hours. Scale bar is 10 μm.

Download video file^{(5.7MB, mp4)}

Movie S9. ER nests can return to peroxisomes during conditions of heightened metabolism, related to Figure 7F–G.

ER nests labelled with SAR1D-mTagBFP2 (blue) initially are dispersed throughout the cytosol in hypocotyl cells of a 4-day-old light-grown seedling but move to and surround peroxisomes (tdT-PTS1; magenta) over time after the seedling is moved to the darkness of the imaging room. Peroxisomes also greatly expand in size during this process. Video shows z-projections (1 μm stack intervals through hypocotyl epidermal cells) acquired every 15 minutes over 12 hours. Scale bar is 10 μm.

Download video file^{(4.9MB, mp4)}

NIHMS2066114-supplement-10.pdf^{(15.8MB, pdf)}

Key resources table

REAGENT or RESOURCE	SOURCE	IDENTIFIER
Antibodies
Mouse anti-HSC70	VWR	Cat# ADI-SPA-817F
Rat anti-HA	Sigma	Cat# 11867423001
Rabbit anti-PMDH	Muhammad et al.¹⁰⁶	N/A
Bacterial and virus strains
Agrobacterium tumefaciens (GV3101)	Koncz and Schell¹⁰³	N/A
NEB 5-alpha E. coli	NEB	Cat# C2987H
Chemicals, peptides, and recombinant proteins
Beta-estradiol	VWR	Cat# AAAL03801-06
Monodansylpentane (MDH)	Abcepta	Cat# SM1000b
Nile red	Sigma	Cat# 72485
LipidTOX deep red	Fisher	Cat# H34477
FM 4-64 ((N-(3-Triethylammoniumpropyl)-4-(6-(4-(Diethylamino) Phenyl) Hexatrienyl) Pyridinium Dibromide))	Invitrogen	Cat# T13320
Oleic acid	Sigma	Cat# O1008-5G
Dioleoylglycerol (DAG)	Sigma	Cat# D8894-50MG
Glyceryl trioleate (TAG)	Sigma	Cat# 92860-10ML
Gibson Assembly Master Mix	NEB	Cat# E2611L
LR Clonase II recombinase	Invitrogen	Cat# 11791-020
Experimental models: Organisms/strains
Arabidopsis thaliana Columbia-0 (Col-0)	Arabidopsis Biological Resource Center	CS60000
Arabidopsis Col-0 (35S:GFP-HDEL)	A gift from Federica Brandizzi; Brandizzi et al.²⁴	N/A
Arabidopsis rhd3-7 (35S:GFP-HDEL)	A gift from Federica Brandizzi; Stefano et al.⁴⁵	SALK 106309 (for rhd3-7)
Arabidopsis pex5-10	Zolman et al.³⁰	SALK_124577
Col-0 (35S-mNG-mPTS^PEX26, UBQ10-mRuby3-PTS1)	Wright and Bartel³	3281; see Figure S1
Col-0 (UBQ10-mRuby3-PTS1)	Wright and Bartel³	3283; see Figure S1
Col-0 (35S-mPTS^PEX22-mNG, UBQ10-mRuby3-PTS1)	Wright and Bartel³	3408; see Figure S1
Col-0 (various transgenes from Figure S1)	This paper	Various;see Figure S1
pex5-10 (4568)	This paper	4568; see Figure S1
rhd3-7 (3408)	This paper	3408; see Figure S1
rhd3-7 (4813)	This paper	4813; see Figure S1
Oligonucleotides
pex5-3 (for pex5-10 wild-type genotyping): GTCGTTGGCTGAATATTTTGTTCGGC	Khan and Zolman³¹	N/A
pex5-21 (for pex5-10 wild-type genotyping): GATATCAAATGCGACTCAAACACTGATGAC	Khan and Zolman³¹	N/A
rhd3-1 (genotyping rhd3-7): GACAGAATATACCGTTACACAGG	This paper	N/A
rhd3-2 (genotyping rhd3-7): GCTGATTAAAGAACAGATGAGAAAG	This paper	N/A
LB1-SALK (for mutant genotyping): CAAACCAGCGTGGACCGCTTGCTGCAACTC	Rasbery et al.³⁸	N/A
Recombinant DNA
See Figure S1 for inserts of plasmids generated and used to transform Col-0 in this study	This paper	N/A
pMCS-GW	Michniewicz et al.¹⁰⁰	N/A
pFZ19	Zhang et al.¹⁰¹	Addgene 36184
pUC57-35S-mTagBFP2-HA, ACT2-TdTomato-PTS1, UBQ10-mNG-mPTS^PEX26	This paper	N/A
pUC57-35S-mTagBFP2-HA, ACT2-TdTomato-PTS1, UBQ10-mPTS^PEX22-mNG	This paper	N/A
Software and algorithms
FIJI	N/A	https://fiji.sc/
Ilastik	Sommer et al.¹⁰⁸	https://www.ilastik.org/
PRISM	N/A	https://www.graphpad.com/
JMP Pro	N/A	https://www.jmp.com/en_us/home.html

Open in a new tab

Highlights.

Peroxisomes and lipid droplets form at specialized ER subdomains, “ER nests”
Lipid-biosynthetic enzymes localize to ER nests
An ER-shaping atlastin and a COPII component localize to and organize ER nests
ER nests can remain associated with progeny organelles and influence their contacts

ACKNOWLEDGMENTS

We thank Matthew Meyer of the Rice University Shared Equipment Authority for electron microscopy, Federica Brandizzi for GFP-HDEL and rhd3-7 seeds, Roxanna Llinas for RNA, and Gabrielle Buck, Isabella Kreko, Roxanna Llinas, Kathryn Smith, and Ana Swearingen for feedback on the manuscript. This work was supported by the National Institutes of Health (R35GM130338 to BB; F31GM150269 to NET) and the Robert A. Welch Foundation (C-1309).

Footnotes

Publisher's Disclaimer: This is a PDF file of an unedited manuscript that has been accepted for publication. As a service to our customers we are providing this early version of the manuscript. The manuscript will undergo copyediting, typesetting, and review of the resulting proof before it is published in its final form. Please note that during the production process errors may be discovered which could affect the content, and all legal disclaimers that apply to the journal pertain.

DECLARATION OF INTERESTS

The authors declare no competing interests.

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Associated Data

This section collects any data citations, data availability statements, or supplementary materials included in this article.

Supplementary Materials

Movie S1. RHD3 localizes in subdomains of ER nests around oleate-induced lipid droplets, related to Figure 5 and S4A–B.

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Movie S2. rhd3^T75A localizes in puncta around lipid droplets, related to Figure 5 and S4C–D.

Download video file^{(3.7MB, mp4)}

Movie S3. SAR1D localizes to ER nest membrane around young peroxisomes, related to Figure 6B.

Download video file^{(2.2MB, mp4)}

Movie S4. ER nests can leave peroxisomes, related to Figure 6C.

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Movie S5. ER nests can shrink after periods of high activity are concluded, related to Figure 6 and Figure S8A.

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Movie S6. ER nests can fuse together, related to Figure 7 and S8B.

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Movie S7. ER nests can divide, related to Figure 7 and S8C.

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Movie S8. ER nests can change their morphology and flatten into sheets, related to Figure 7 and S8D.

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Movie S9. ER nests can return to peroxisomes during conditions of heightened metabolism, related to Figure 7F–G.

Download video file^{(4.9MB, mp4)}

NIHMS2066114-supplement-10.pdf^{(15.8MB, pdf)}

Data Availability Statement

The data supporting the findings are in the main text or supplementary information.
This paper does not report original code.
Any additional information required to reanalyze the data reported in this paper is available from the lead contact upon request.

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PERMALINK

ER nests are specialized ER subdomains in Arabidopsis where peroxisomes and lipid droplets form

Zachary J Wright

Nathan E Tharp

Bonnie Bartel

SUMMARY

Graphical Abstract

eTOC blurb

INTRODUCTION

RESULTS

De novo peroxisome formation is active and can be observed in Arabidopsis seedlings

Figure 1. De novo peroxisome formation is active in Arabidopsis seedlings and occurs at complex ER domains – ER nests.

De novo peroxisome formation occurs at discrete ER domains—ER nests

Lipid droplets can form from ER nests

Figure 2. Lipid droplets can form from ER nests.

Lumenal protein import drives peroxisome expansion in ER nests

Figure 3. Lumenal protein import drives peroxisome expansion in ER nests.

Sterol biosynthetic enzymes localize to ER nests, and nest membrane remains with mature lipid droplets

Figure 4. Sterol biosynthetic enzymes localize to ER nests, and nest membrane remains associated with mature lipid droplets.

RHD3/atlastin functions in ER nest organization

Figure 5. RHD3/atlastin functions in ER nest organization.

ER nests harbor SAR1 isoforms

Figure 6. ER nests harbor SAR1 isoforms.

ER nests remaining around peroxisomes appear to control ILV formation

Figure 7. ER nests restrict ILV formation sites and define peroxisome–lipid droplet contacts.

ER nests are dynamic and can re-enclose organelles during heightened metabolism

DISCUSSION

Limitations of the study

RESOURCE AVAILABILITY

Lead contact

Materials availability

Data and code availability

STAR METHODS

EXPERIMENTAL MODEL AND STUDY PARTICIPANT DETAILS

Arabidopsis

METHOD DETAILS

DNA cloning methods

Arabidopsis transformation

Estradiol induction

Lipid droplet induction

Immunoblotting

Confocal microscopy

Transmission electron microscopy

Lipid analysis

QUANTIFICATION AND STATISTICAL ANALYSIS

Supplementary Material

Highlights.

ACKNOWLEDGMENTS

Footnotes

REFERENCES

Associated Data

Supplementary Materials

Data Availability Statement

ACTIONS

PERMALINK

RESOURCES

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