Regulation of synaptic development and function by the Drosophila PDZ protein Dyschronic
ABSTRACT Synaptic scaffold proteins control the localization of ion channels and receptors, and facilitate molecular associations between signaling components that modulate synaptic transmission and plasticity. Here, we define novel roles for a recently described scaffold protein, Dsychronic (DYSC), at the Drosophila larval neuromuscular junction. DYSC is the Drosophila homolog of whirlin/DFNB31, a PDZ domain protein linked to Usher syndrome, the most common form of human deaf-blindness. We show that DYSC is expressed presynaptically and is often localized adjacent to the active zone, the site of neurotransmitter release. Loss of DYSC results in marked alterations in synaptic morphology and cytoskeletal organization. Moreover, active zones are frequently enlarged and misshapen in dysc mutants. Electrophysiological analyses further demonstrate that dysc mutants exhibit substantial increases in both evoked and spontaneous synaptic transmission. We have previously shown that DYSC binds to and regulates the expression of the Slowpoke (SLO) BK potassium channel. Consistent with this, slo mutant larvae exhibit similar alterations in synapse morphology, active zone size and neurotransmission, and simultaneous loss of dysc and slo does not enhance these phenotypes, suggesting that dysc and slo act in a common genetic pathway to modulate synaptic development and output. Our data expand our understanding of the neuronal functions of DYSC and uncover non-canonical roles for the SLO potassium channel at Drosophila synapses. KEY WORDS: PDZ domain, Scaffold protein, Active zone, BK channel, Drosophila, Neuromuscular junction
Elucidating the molecular mechanisms of synapse development and function is important for understanding biological processes such as learning and memory, as well as neurological diseases such as autism spectrum disorder and schizophrenia (Caroni et al., 2012; Sudhof, 2008; Wondolowski and Dickman, 2013). Crucial to correct synaptic organization are scaffold proteins, which act as organizing hubs that link ion channels, neurotransmitter receptors and other membrane proteins to the cytoskeleton and to downstream intracellular signaling networks (Emes and Grant, 2012; Verpelli et al., 2012). In a screen for Drosophila with altered circadian
Department of Neuroscience, The Farber Institute for Neurosciences, Thomas 2 Jefferson University, Philadelphia, PA 19107, USA. UCL Institute of Neurology, 3 London WC1N 3BG, UK. Department of Neurology, John Hopkins University, Baltimore, MD 21287, USA. *These authors contributed equally to this work ‡ These authors contributed equally to this work §
Received 26 February 2014; Accepted 16 September 2014
behavior, we recently identified a previously uncharacterized gene encoding a scaffold protein with multiple PDZ domains that we termed dyschronic (dysc). Loss of DYSC results in arrhythmic patterns of circadian locomotor activity, and dysc is the Drosophila homolog of the human locus whirlin/DFNB31, mutations in which lead to Usher syndrome (USH), a leading cause of deaf-blindness (Jepson et al., 2012; Mburu et al., 2003). We found that DYSC binds to and controls the expression of the Slowpoke (SLO) BK potassium channel in the adult Drosophila nervous system, thus identifying DYSC as a novel ion channel regulator that impacts complex behavior (Jepson et al., 2012). BK potassium channels are synergistically activated by membrane depolarization and elevated intracellular Ca2+ (Barrett et al., 1982; Lee and Cui, 2010; Prakriya and Lingle, 1999). In neurons, BK channels limit action potential duration by linking Ca2+ influx through voltage-gated Ca2+ channels (VGCCs) to potassium efflux (Bean, 2007). Interestingly, in addition to the axon, BK channels are also localized at the synapse near to the active zone (AZ) (Hu et al., 2001), a crucial structural determinant of the probability of vesicle fusion (Sigrist and Schmitz, 2011). In Drosophila, the AZ is an electron-dense macromolecular structure and functions in part to tether synaptic vesicles in close proximity to VGCCs (Atwood et al., 1993; Koenig and Ikeda, 1999); recent genetic studies have uncovered a number of core constituents of the Drosophila AZ as well as proteins involved in its assembly (Johnson et al., 2009; Kaufmann et al., 2002; Kittel et al., 2006; Liu et al., 2011; Nieratschker et al., 2009; Wagh et al., 2006). For example, Bruchpilot (BRP) is a key component of T-bars, dense bodies that project intracellularly from the AZ core and connect to a subpopulation of synaptic vesicles (Atwood et al., 1993; Koenig and Ikeda, 1999; Reist et al., 1998; Wagh et al., 2006). Using super-resolution microscopy, BRP complexes have been shown to form donut-like rings around a central core of VGCCs, and loss of BRP results in reduced VGCC localization in the AZ and a substantial decrease in evoked excitatory currents (Kittel et al., 2006). However, whereas much research has focused on molecules that promote AZ formation and neurotransmitter release, less is known regarding factors that negatively regulate the AZ and vesicle fusion. Here, we demonstrate that DYSC and SLO coordinately influence diverse aspects of synaptic development and function at the Drosophila larval neuromuscular junction (NMJ), and act as negative regulators of AZ size and evoked neurotransmission. Most surprisingly, spontaneous vesicle fusion is enhanced in dysc and slo mutants, even in zero-Ca2+ conditions, suggesting that SLO channels impact spontaneous neurotransmission independently of enhanced VGCC activity. Our results reveal intriguing noncanonical roles for the SLO BK channel and significantly expand the known functions of the USH homolog DYSC in the Drosophila nervous system. 1
James E. C. Jepson1,2, *, Mohammed Shahidullah1,*, Die Liu1,‡, Sylvain J. le Marchand1,‡, Sha Liu3, Mark N. Wu3, Irwin B. Levitan1, Matthew B. Dalva1 and Kyunghee Koh1,§
Development (2014) 141, 1-10 doi:10.1242/dev.109538
RESULTS DYSC regulates synaptic morphology and cytoskeletal organization
Fig. 1. DYSC regulates synaptic morphology and Futsch localization. (A) Schematic of the dysc transcription unit, locations of the dysc c03838 and dysc s168 P-element insertions and the MiMIC protein trap insertion used to generate the dysc::gfp allele. Genomic regions encoding the three PDZ domains are also shown. (B) Confocal projections showing representative NMJs at muscle 6/7, segment 3. Neuronal membranes are labeled with antiHRP. Arrowheads indicate enlarged synaptic boutons in dysc s168 and dysc c03838 homozygotes that are absent in controls or heterozygotes. Scale bar: 20 μm. (C,D) Average number of total synaptic boutons (C) and average bouton size (D) in wild-type controls, and in dysc s168 and dysc c03838 heterozygotes and homozygotes, and dysc c03838/dysc s168 transheterozygotes. (C) n=12-29 synapses. (D) n=167-211 boutons. (E) Confocal slices illustrating localization of Futsch in synaptic termini in wild-type controls and dysc s168 and dysc c03838 homozygotes. Arrowheads indicate synaptic boutons exhibiting unusual unbundled accumulation of Futsch. Scale bar: 5 μm. (F) Quantification of the mean proportion of boutons exhibiting unbundled Futsch. n=14-16 synapses, 265-381 boutons. Values represent mean±s.e.m. ***P<0.0005; ns, not significant (P>0.05), one-way ANOVA with Dunnett post-hoc test.
DYSC localizes to synaptic termini and is required presynaptically
We next sought to examine the localization of DYSC at the larval NMJ. Immunohistochemical experiments using our previously described anti-DYSC antibody failed to generate robust signals at larval synapses, possibly due to low efficacy and/or low DYSC expression. To make use of a higher affinity antibody, we used the MiMIC system (Venken et al., 2011) to insert a GFP/FlASH/ StrepII/3xFLAG protein trap upstream of the region encoding the
We have previously found that DYSC is broadly localized within the adult fly brain and exhibits strong expression in axonal tracts throughout the nervous system (Jepson et al., 2012). However, DYSC is also present in synaptic neuropil regions (Jepson et al., 2012). We were therefore intrigued by the possibility that DYSC might play important roles at Drosophila synapses. To test whether DYSC regulates synaptic development and/or function, we turned to the NMJ of third instar Drosophila larvae, a model glutamatergic synapse, the structural and functional properties of which have been extensively studied. We initially examined synaptic morphology of dysc mutants using two transposon insertion alleles (dysc c03838 and dysc s168) (Fig. 1A). The dysc transcription unit generates protein isoforms containing two or three PDZ domains (Jepson et al., 2012) (Fig. 1A; www. flybase.org). Both long and short DYSC isoforms are undetectable by western blotting of dysc c03838 head extracts (Jepson et al., 2012), and dysc cDNA expression was not detected in dysc c03838 head tissue (supplementary material Fig. S1A), indicating that dysc c03838 is a null allele. By contrast, the dysc s168 insertion selectively disrupts expression of long DYSC isoforms containing three PDZ domains, while leaving the short isoform intact (Jepson et al., 2012). Both dysc c03838 and dyscs168 homozygotes exhibited a ∼40% reduction in the total number of synaptic boutons at muscle 6/7 (Fig. 1B,C), resulting from a decrease in the number of both type 1b and type 1s boutons (supplementary material Fig. S1B,C). A similar phenotype was observed in dysc c03838/dysc s168 trans-heterozygotes, but there was no significant alteration in bouton number in dysc c03838/+ or dyscs168/+ heterozygotes (Fig. 1B,C; supplementary material Fig. S1B,C). dysc homozygous and trans-heterozygous mutants also exhibited a substantial increase in the average size of type 1b boutons (Fig. 1B,D). By contrast, mean synaptic area and branch number were not altered in dysc mutants (supplementary material Fig. S1D,E). Similar phenotypes in terms of bouton number and size were observed in larval synapses innervating muscle 4 (supplementary material Fig. S1F,G). Thus, DYSC regulates the number and size of larval synaptic boutons but not overall synaptic size. Because whirlin, the mammalian DYSC homolog, has recently been shown to affect the stability of the axonal cytoskeleton (Green et al., 2013), we wondered whether DYSC modulates synaptic morphology by controlling the localization of synaptic cytoskeletal proteins. We therefore visualized expression of the microtubule-binding protein Futsch in wild-type and dysc mutant synapses (Fig. 1E). Futsch-mediated regulation of microtubule organization is important for correct synaptic bouton budding (Roos et al., 2000), and mislocalization of synaptic Futsch in various Drosophila mutants has been associated with reduced synaptic bouton number and increased bouton size (Mosca et al., 2012; Pennetta et al., 2002). Consistent with previous data (Roos et al., 2000), filamentous ‘tendrils’ and structured ‘loops’ of Futsch-bound microtubules were generally observed in wild-type synapses (Fig. 1E). By contrast, ∼25% of dysc mutant synapses exhibited an abnormal mode of Futsch localization, characterized by a substantially increased level of unstructured Futsch within synaptic boutons (Fig. 1E,F; supplementary material Fig. S2). This pattern of Futsch expression was rarely observed in wild-type synaptic boutons (∼2.6%, Fig. 1F). Thus, loss of DYSC leads to an alteration in the synaptic localization of a key cytoskeletal regulator in a manner consistent with an inhibition of synaptic bouton budding in dysc mutants.
third PDZ domain (Fig. 1A). This strategy allowed labeling of DYSC isoforms containing three PDZ domains using anti-GFP and anti-FLAG antibodies. We term this new dysc allele dysc::gfp. The protein trap exon does not disrupt the PDZ domain-coding sequences within the dysc locus, and importantly, dysc::gfp larvae exhibited normal synaptic morphology (supplementary material Fig. S3A-C). Furthermore, dysc::gfp adult flies exhibited rhythmic patterns of circadian locomotion under constant-dark conditions (data not shown). Thus, the GFP insertion does not inhibit the normal function of DYSC. In both the adult and the larval Drosophila nervous systems, GFPtagged DYSC was localized to axonal regions and, importantly, to the synaptic neuropil, as evidenced by colocalization with the presynaptic marker Bruchpilot (BRP) (Kittel et al., 2006; Wagh et al., 2006) (supplementary material Fig. S3D,E). At the larval NMJ, full-length DYSC detected using an anti-GFP antibody exhibited a punctate pattern of localization in both motor neuron axon bundles and in synaptic bouton termini (Fig. 2A). A similar expression pattern was observed when using an anti-FLAG antibody to detect DYSC (supplementary material Fig. S4A), and this pattern was not observed in larvae lacking the dysc::gfp allele using either anti-GFP (Fig. 2A) or anti-FLAG antibodies (supplementary material Fig. S4A). Furthermore, neuronal overexpression of a long DYSC isoform in combination with an anti-DYSC antibody generated similar presynaptic punctate signals (supplementary material Fig. S4B,C). The localization of DYSC::GFP suggested a presynaptic function for DYSC. Consistent with this hypothesis, pre- but not postsynaptic expression of a full-length DYSC transgene (uas-dysc-long)
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significantly rescued both the reduced bouton number and increased bouton size of dysc mutants (Fig. 2B-D). Transgenic restoration of DYSC solely in neurons also rescued the mislocalization of synaptic Futsch (supplementary material Fig. S5). The similar phenotypes of dysc c03838 and dysc s168 homozygotes further suggested that full-length DYSC isoforms containing three PDZ domains are specifically required for normal synaptic bouton proliferation, as expression of the short DYSC isoform containing the two N-terminal PDZ domains is maintained at wild-type levels in dysc s168 homozygotes (Jepson et al., 2012). We therefore performed rescue experiments using a transgene encoding a short DYSC isoform (uas-dysc-short), and indeed found that neither neuronal nor muscle expression of the short DYSC isoform rescued any aspect of synaptic morphology (Fig. 2C,D). In addition, to test whether DYSC levels influence synaptic bouton size and number in a dose-dependent manner, we overexpressed DYSC in neurons in a wild-type background. Overexpression of either long or short DYSC transgenes did not alter bouton number or size relative to controls (supplementary material Fig. S6). These experiments indicate that DYSC is required in the presynaptic compartment to influence synaptic development, and suggest that the C-terminal region that includes the third PDZ domain of DYSC is required for this function. DYSC influences the architecture of active zones
We next sought to examine the subcellular localization of presynaptic DYSC puncta in more detail by immunostaining synapses in dysc::gfp larvae with anti-GFP and either antiBruchpilot (anti-BRP) or anti-Fasciclin II (FASII) antibodies.
Fig. 2. DYSC is required in neurons for correct synaptic development. (A) Confocal slices illustrating punctate expression of DYSC::GFP (GFP) in both axonal bundles descending from the larval ventral nerve cord and at synaptic bouton termini. This staining pattern was not observed in controls lacking the dysc::gfp insertion. Scale bars: 20 μm in axons; 5 μm in boutons. (B) Confocal projections showing representative NMJs at muscle 6/7, segment 3 of wild-type controls, dysc c03838 (dysc) homozygotes and dysc larvae with presynaptic expression of a uas-dysc-long transgene. Neuronal membranes are labeled with antiHRP. Arrowheads indicate enlarged synaptic boutons in dysc larvae that are absent following presynaptic restoration of full-length DYSC. Scale bar: 20 μm. (C,D) Pre- and postsynaptic rescue experiments examining bouton number (C; n=12-20) or bouton size (D; n=176-326) using either elav-Gal4 (neurons) or 24B-Gal4 (muscle) drivers to express short or long DYSC isoforms. For ease of viewing, the data for elav-Gal4/+ and 24B-Gal4/+ synapses are repeated on each side of the graph. Values represent mean±s.e.m. ***P<0.0005; ns, not significant (P>0.05), one-way ANOVA with Tukey’s post-hoc test.
BRP is a key constituent of T-bars, electron-dense protrusions found at the AZ (Kittel et al., 2006; Wagh et al., 2006), whereas FASII is a marker for periactive zones linked to endocytosis (Sone et al., 2000). DYSC::GFP puncta were often (but not always) adjacent to or overlapping with BRP-labeled T-bars (Fig. 3A, upper panel). Transgenic full-length DYSC expressed presynaptically exhibited a similar localization near BRP puncta (supplementary material Fig. S4C). By contrast, DYSC::GFP exhibited little overlap with FASII (Fig. 3A, lower panel), indicating that synaptic DYSC is preferentially localized in close proximity to AZs. Given the close association of DYSC and BRP-labeled T-bars, we next investigated whether DYSC influences the expression or localization of BRP. Under confocal microscopy, we observed no clear alteration either in the localization of BRP or the apposition between BRP and the postsynaptic Glutamate Receptor IIC (GLURIIC) in dysc mutant synapses (supplementary material Fig. S7A,B), nor any significant change in GLURIIC levels (supplementary material Fig. S7B,C). However, at increased
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magnification, the size of BRP puncta appeared to be somewhat increased in dysc mutants (supplementary material Fig. S7B). These observations suggested that DYSC might impact the fine structure of T-bars. We therefore used a super-resolution imaging technique, stimulated emission depletion (STED) microscopy (Dyba et al., 2003), to obtain nanometer-scale images of BRP structures wild-type and dysc mutant synapses (Fig. 3B). As shown previously (Kittel et al., 2006; Liu et al., 2011; Owald et al., 2012), BRP forms donut-like structures in wild-type synapses when observed under STED imaging (Fig. 3B). Intriguingly, we found that both dysc c03838 homozygous and dysc c03838/dysc s168 transheterozygous mutants exhibited a significant increase in the mean area of individual BRP rings (Fig. 3B,C). Furthermore, we observed frequent merging of neighboring BRP rings in dysc mutants (Fig. 3B). These phenotypes were rescued by neuronal expression of a full-length DYSC transgene, confirming that this phenotype is caused by a loss of presynaptic DSYC (Fig. 3D,E). Thus, in addition to modulating synaptic morphology, DYSC also plays a significant role in regulating the macromolecular architecture of a key constituent of the AZ.
Fig. 3. DYSC modulates the structural composition of active zones. (A) Confocal slices illustrating subcellular localization of DYSC::GFP (GFP) in the presynaptic compartment. DYSC::GFP puncta are often closely associated with the AZ component Bruchpilot (BRP, upper panel). By contrast, there is little overlap between DYSC::GFP and the periactive zone marker FASII (lower panel). Scale bar: 5 μm. (B) Representative confocal (top) and STED (bottom) slices of BRP-marked AZs in control larvae and two dysc allelic backgrounds. Arrowheads indicate clustered AZs. Scale bar: 1 μm. (C) Average area for individual AZs in control and dysc mutant synapses. n=39-56 AZs. (D) Representative STED images of AZ organization in dysc c03838 (dysc) mutants harboring the pan-neuronal elav-Gal4 (elav) driver, a uas-dysc-long transgene (dysc-long) or both. Arrowheads indicate clustered AZs. Scale bar: 1 μm. (E) Average area of individual AZs in the above genotypes. Values represent mean±s.e.m. n=29-57 AZs. **P<0.005; ***P<0.0005, one-way ANOVA with Dunnett post-hoc test.
We have previously shown that DYSC binds to and promotes the expression of SLO channels in the adult Drosophila nervous system (Jepson et al., 2012). However, whether this is also the case at earlier developmental time points is unclear. Consistent with our previous data in the adult brain, loss of DYSC also resulted in a robust reduction in SLO expression in motor neuron axons of the larval ventral nerve cord (supplementary material Fig. S8A). Previous work has shown that mutations in slo lead to an increased frequency of rhythmic peristalsis of the body wall muscle (McKiernan, 2013), and, similarly, we found that larvae homozygous for both dysc s168 and dysc c03838 also exhibited a significant enhancement in the rate of peristalsis (supplementary material Fig. S8B). We next asked whether the alterations in synaptic bouton morphology found in dysc mutants were also observed in larvae lacking SLO channels. To this end, we obtained a slo mutant line carrying a Minos element insertion in the exon encoding transmembrane domain 2 of the SLO channel (slo 11481; www. flybase.org), resulting in an interruption in the coding sequence that is expected to inhibit production of full-length slo transcripts, and examined how this mutation affects synaptic development. We confirmed that, as predicted, slo 11481 is a null allele using RT-PCR (supplementary material Fig. S9A). In addition, to control for differences in genetic background, we outcrossed all dysc and slo alleles into an isogenic white 1118 (iso31) background for 5-10 generations. Like dysc mutants, slo 11481 larval synapses exhibited significantly increased bouton areas (Fig. 4A,C). This phenotype was also observed in genetic backgrounds containing trans-heterozygotic combinations of slo 11481, slo 4 (a canonical slo null allele resulting from a large chromosomal inversion), slo 1 (a null or strongly hypomorphic allele) and slo Df (a deficiency removing the slo locus) (supplementary material Fig. S9B,D,F), and was rescued by the introduction of a slo transgene (Fig. 4C) (Ghezzi et al., 2004). Interestingly, in contrast to dysc mutants, synaptic bouton number was not significantly altered in any of the slo mutants we examined (Fig. 4B and supplementary material Fig. S9C,E). This contrasts with previous work reporting that larvae homo- or transheterozygous for three slo alleles (slo 1, slo 98 and slo 4) exhibit a hyper-budding phenotype at larval synapses (Lee and Wu, 2010). Our experiments
SLO channels impact synaptic development at the larval NMJ
Development (2014) 141, 1-10 doi:10.1242/dev.109538
Fig. 4. slo and dysc act in the same pathway to regulate synaptic bouton morphology. (A) Representative confocal projections of synapses from control larvae, slo 11481 heterozygotes and homozygotes, dysc s168 homozygotes, and dysc s168, slo 11481 double mutants labeled with anti-HRP. Arrowheads indicate enlarged boutons that are present in slo 11481 and dysc s168 homozygotes, as well as dysc s168, slo 11481 double mutants. Scale bar: 20 μm. (B,C) Average bouton number (B) and bouton area (C) in wild-type controls, slo 11481 heterozygotes, slo and dysc homozygotes, dysc, slo double mutants, and slo 11481 homozygotes harboring a slo rescue construct (B52H). n=10-40 synapses and 195-463 boutons. Values represent mean±s.e.m. *P<0.05; ***P<0.0005; ns, not significant (P>0.05), one-way ANOVA with Tukey’s post-hoc test.
dysc and slo act in a common genetic pathway to regulate synaptic bouton morphology
To examine whether slo and dysc are components of the same genetic pathway, we generated slo, dysc double mutants using two independent alleles of dysc and the slo 11481 allele, and compared their synaptic phenotype relative to individual slo and dysc mutations. In relation to bouton size, slo, dysc double mutants exhibited significantly increased bouton size relative to controls (P<0.0001), but were not significantly different when compared with dysc mutations alone (P>0.99, one-way ANOVA with Tukey’s post-hoc test) (Fig. 4B,D). Furthermore, we observed a significant increase in bouton size in slo 11481/dysc c03838 transheterozygotes relative to controls and heterozygotes for either allele (supplementary material Fig. S9G), demonstrating a synergistic interaction between the two genes. Unexpectedly, we found that loss of SLO suppressed the reduction in synaptic bouton number observed in dysc mutants (Fig. 4B,C). A number of mechanisms could account for this effect. For example, loss of DYSC may lead to an altered subcellular localization of SLO that results in reduced bouton number, which is rescued by complete loss of SLO. Alternatively, selective loss of SLO in neurons but not in muscle tissue in dysc mutants may affect bouton number, whereas complete loss of SLO does not. We were
unable to test these competing hypotheses because synaptic SLO could not be detected using our previously described antibody (Jepson et al., 2012) or a SLO::GFP fusion generated using the MiMIC system (Venken et al., 2011) (data not shown). Nonetheless, the genetic interactions between dysc and slo alleles detailed above strongly suggest that DYSC and SLO act in a common pathway to influence synaptic bouton size and number. SLO channels impact active zone structure and Futsch localization
As dysc mutants display alterations in the structure of a key AZ complex, we determined whether slo mutants also exhibit similar AZ defects. Under STED microscopy, larvae homozygous for the slo 11481 allele indeed exhibited an increase in BRP ring area and clustering that was almost identical to that seen in dysc larvae (Fig. 5). Larvae lacking a distinct potassium channel, Shaker, did not exhibit either an increase in BRP area or merging of BRP rings, suggesting that the alterations in AZ structure in slo mutants are not due to a general effect of neuronal hyperexcitability (supplementary material Fig. S11). To test whether SLO and DYSC act in the same pathway for the regulation of AZ morphology, we visualized BRP rings in dysc, slo double mutants. The phenotype of dysc, slo double mutants was statistically indistinguishable from each individual mutation (Fig. 5A,B; P>0.27, one-way ANOVA with Tukey’s post-hoc test), suggesting that DYSC and SLO act in the same pathway to regulate AZ size. Furthermore, we also observed a mislocalization of synaptic Futsch in slo mutants similar to that observed following loss of DYSC (supplementary material Fig. S12). Collectively, the above data provide evidence that DYSC and SLO act in a common pathway to regulate synaptic morphology, cytoskeletal dynamics and presynaptic macromolecular organization. DYSC and SLO inhibit evoked neurotransmission
To test whether DYSC also impacts the physiology of larval synapses, we recorded evoked excitatory junctional currents (EJCs) in muscle 6/7 under voltage-clamp mode. We observed a significant increase in both the EJC amplitude and width in dysc c03838 larvae relative to controls (Fig. 6A-C). The enhancement of the EJC amplitude was rescued by presynaptic expression of a full-length DYSC transgene in dysc c03838 homozygotes (Fig. 6D,E), and there 5
were performed at 25°C in an iso31 genetic background, whereas the previous analysis was performed at room temperature in a Canton-S background (Lee and Wu, 2010). Phenotypic effects of mutations may be modified by both environmental conditions and genetic background (Chari and Dworkin, 2013; Dworkin et al., 2009; Wang et al., 2014). Indeed, of the three slo mutants we examined raised at a lower temperature of 18°C (slo 11481 and slo 4 homozygotes, and slo 1/slo 4 transheterozygotes), only slo 4 homozygotes exhibited a measurable increase in synaptic bouton budding (supplementary material Fig. S10). The slo 4 chromosome contains a large inversion that prevents recombination, and it is likely that despite being outcrossed to an iso31 background five times, slo 4 retains a substantial amount of the original background. Thus, slo 4 mutants may harbor additional mutations that impact synaptic development in a temperature-sensitive manner. We conclude that in an iso31 genetic background, SLO channels predominantly act to negatively regulate bouton size rather than suppressing synaptic bouton budding.
Fig. 5. DYSC and SLO act in concert to regulate AZ structure. (A) Representative STED slices of BRP-labeled AZs in wild-type control, individual slo 11481 and dysc c03838 homozygotes, and dysc c03838, slo 11481 double mutant larval synapses. Scale bar: 1 μm. Arrowheads indicate clustered AZs. (B) Average BRP ring area for the above genotypes. Values represent mean±s.e.m. n=40-52 AZs; ns, not significant (P>0.05), ***P<0.0005, one-way ANOVA with Tukey’s post-hoc test.
was a trend towards a rescue of the increased EJC width in the same genetic background (Fig. 6F). Thus, presynaptic DYSC negatively regulates evoked synaptic transmission. The larger and broader action potentials in dysc mutants would be predicted to result in higher concentrations of intracellular Ca2+. As the levels of intracellular Ca2+ near the sites of neurotransmitter release influence the probability of vesicle fusion (Katz and Miledi, 1963), we examined whether dysc mutants also exhibited alterations in short-term synaptic plasticity using a paired-pulse protocol. Consistent with previous data (Kittel et al., 2006), paired EJCs from wild-type NMJs were approximately constant over an interval of 30-130 ms (supplementary material Fig. S13). By contrast, EJCs from dysc mutant NMJs exhibited significant paired-pulse facilitation over this interval (supplementary material Fig. S13). Thus, in addition to altering the basal properties of evoked currents, DYSC also regulates short-term plasticity.
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As DYSC promotes neuronal SLO channel expression (Jepson et al., 2012), we also tested whether DYSC and SLO act in the same genetic pathway to regulate evoked synaptic transmission by examining evoked currents in slo and dysc, slo double mutants (Fig. 6A-C). Similar, to dysc mutants, slo mutants exhibited a significant increase in EJC amplitude and width (Fig. 6A-C). This contrasts with previous studies that reported no significant alterations of EJC amplitudes in slo mutants (Lee et al., 2008, 2014), but these discrepancies may due to different genetic backgrounds as discussed above. Importantly, dysc, slo double mutants also exhibited increased EJC amplitude and width, and did not show enhanced phenotypes over single mutants (Fig. 6A-C), suggesting that DYSC and SLO act within the same pathway to negatively control evoked synaptic output. DYSC and SLO inhibit spontaneous neurotransmitter release via a Ca2+-independent mechanism
We next asked whether DYSC also modulates spontaneous vesicle fusion by examining miniature excitatory junctional currents (mEJCs) from control and dysc c03838 homozygotes (Fig. 7A). We observed a striking (∼250%) increase in the frequency of mEJCs in dysc mutants (Fig. 7A,B), indicating that loss of DYSC results in a substantial enhancement of the probability of spontaneous vesicle release. We also observed a similar increase in mEJC frequency in slo11481 homozygotes, and analysis of mEJCs in dysc, slo double mutants demonstrated that the effects of these mutations were non-additive, as there were no significant differences in mEJC frequency in any of the mutant backgrounds examined (P>0.9, Kruskal-Wallis test with Dunn’s post-hoc test) (Fig. 7B). In addition, we observed a trend towards reduced mEJC amplitudes in dysc and slo homozygotes, as well as dysc, slo double mutants, but this trend only achieved significance in slo mutants (Fig. 7C). Thus, with regard to spontaneous neurotransmission, DYSC and SLO primarily act in a common pathway to regulate the frequency of synaptic vesicle fusion. As both dysc and slo mutants exhibit structural alterations in AZ architecture, one simple hypothesis is that the larger (and merged)
Fig. 6. DYSC and SLO inhibit evoked synaptic transmission. (A) Representative evoked excitatory junctional current (EJC) traces in wild-type control larvae, dysc and slo homozygotes, and dysc, slo double mutants. EJCs were measured from muscle 6/7. (B,C) Quantification of mean EJC amplitude (B) and width at 50% rise (C) in the above genotypes. n=7-21. (D-F) Representative EJC traces (D) and quantification of mean EJC amplitude (E) and width (F) from dysc mutants carrying a full-length dysc transgene (uas-dysclong) alone or in combination with the pan-neuronal elav-Gal4 driver (elav). n=6-15. Data are presented as modified box plots showing the median, 25th and 75th percentiles. ns, not significant (P>0.05), *P<0.05, **P<0.005, ***P<0.0005, Kruskal-Wallis test with Dunn’s post-hoc test (B,C) or MannWhitney U-test (E,F).
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AZs in dysc and slo mutants facilitate enhanced spontaneous vesicle fusion. The BRP ring-like complex within the AZ surrounds a central core of VGCCs (Kittel et al., 2006). Interestingly, recent work in mammalian synapses has shown that stochastic opening of VGCCs in the AZ is responsible for a substantial proportion of spontaneous neurotransmitter release (Ermolyuk et al., 2013), and that AZ size positively correlates with the frequency of spontaneous neurotransmitter release (Holderith et al., 2012; Matz et al., 2010), presumably via altered VGCC occupancy. If enhanced spontaneous vesicle fusion in dysc and slo mutants was due to an increase in the number of VGCCs within the AZ, this effect should be suppressed by removing extracellular Ca2+. However, recording mEJCs in zero-Ca2+ saline did not alter mEJC frequency or amplitude in either control or mutant backgrounds (Fig. 7B,C). Thus, the effect of DYSC and SLO on spontaneous release is independent of voltage-gated Ca2+ channels and may instead be directly due to structural alterations at AZs that facilitate an increased probability of synaptic vesicle fusion. DISCUSSION
Our work has revealed novel functions for the PDZ domain protein DYSC and its binding partner SLO in modulating diverse aspects of synaptic development and physiology, including synaptic bouton size, cytoskeletal protein localization, AZ structure and synaptic transmission. With respect to synaptic physiology, we found that DYSC acts as a negative regulator of evoked neurotransmission as well as of short-term synaptic plasticity; the alteration in evoked currents in dysc mutants are consistent with the known function of DYSC as a positive regulator of SLO channels (Jepson et al., 2012). Surprisingly, in addition to inhibiting evoked current amplitude and duration, DYSC and SLO also negatively regulate spontaneous neurotransmitter release in a manner that is independent of VGCC channel activation, as increased rates of miniature events are still robustly detected under zero-Ca2+ conditions. This finding raises the intriguing possibility that SLO channels perform non-canonical structural roles that impact spontaneous neurotransmission, as well as synaptic morphology and AZ architecture.
One issue emerging from the above results is whether altered AZ structure and spontaneous neurotransmission in dysc and slo mutants arise via a homeostatic response to increased excitability. Loss of specific ion channels in Drosophila has been shown to activate homeostatic transcriptional networks that act to balance excitability (Parrish et al., 2014; Ping and Tsunoda, 2012). In addition, the AZ T-bar itself has been theorized to function as a ‘plasticity module’ that can undergo structural adjustments in response to changes in synaptic input that, in turn, modify the probability of vesicle release (Wichmann and Sigrist, 2010). For example, at the crayfish NMJ, electrophysiological stimulation results in an increased number of AZs containing multiple T-bars (Wojtowicz et al., 1994), similar to the merged BRP rings observed in dysc and slo mutants. However, although we cannot rule out such a mechanism, we argue for a more direct role of the DYSC/ SLO complex in regulating AZ formation and spontaneous neurotransmission. Numerous studies have shown that reducing postsynaptic excitability at the larval NMJ results in a compensatory increase in the probability of neurotransmitter release via a retrograde signaling mechanism (Frank et al., 2006; Muller and Davis, 2012). By contrast, in dysc and slo mutants both evoked and spontaneous neurotransmission are enhanced (Figs 6 and 7), a finding that is inconsistent with a homeostatic negativefeedback model. Given the close proximity of DYSC to synaptic T-bars, we suggest instead that the DYSC/SLO complex directly controls AZ structure in a manner that leads to increased spontaneous vesicle fusion. Such an effect has been demonstrated for Drosophila Neurexin 1 (NRX-1), a synaptic cell-adhesion molecule that is also localized close to T-bars (Li et al., 2007; Owald et al., 2012). Similar to DYSC and SLO, NRX-1 negatively regulates BRP ring size and inhibits merging of BRP rings (Owald et al., 2012); these structural alterations correlate with an increased mEJP frequency in nrx-1 mutants (Li et al., 2007). How the changes in AZ architecture observed in dysc, slo and nrx-1 mutants are linked to enhanced rates of vesicle fusion is unclear. One possibility is that enlarged and misshapen AZs are less efficient at clamping synaptic vesicles to prevent premature exocytosis in the absence of Ca2+ (Huntwork and Littleton, 2007). 7
Fig. 7. DYSC and SLO inhibit spontaneous synaptic transmission independently of extracellular Ca2+. (A) Representative mEJC traces illustrating spontaneous neurotransmitter release from larval presynaptic terminals in wildtype control larvae, dysc and slo homozygotes, and dysc, slo double mutants. (B,C) Mean mEJC frequency (B) and amplitude (C) in 1 mM (+) or 0 mM (−) extracellular Ca2+. n=5-8 per genotype, with each data point representing the average of an individual trace. Data are presented as modified box plots showing the median, 25th and 75th percentiles. ns, not significant (P>0.05), *P<0.05, **P<0.005, Kruskal-Wallis test with Dunn’s post-hoc test.
MATERIALS AND METHODS Drosophila genetics and molecular biology
Flies were reared on standard food containing cornmeal, yeast and molasses at 25°C under 12 h light:12 h dark cycles. The dysc c03838 and dysc s168 P-element insertions have been described previously (Jepson et al., 2012). The elav-Gal4 and 24B-Gal4 stocks, flies harboring the slo 11481 Minos element, a deficiency line that removes the slo locus [w 1118; +; Df(3R)BSC397] and the dysc Mi02729 MiMIC insertion used to generate the dysc::gfp allele were obtained from the Bloomington Stock Center. To control for potential differences in genetic background, all dysc and slo alleles were outcrossed into a white 1118 (iso31) background for 5-10 generations prior to experimentation. The dysc::gfp allele was generated by ΦC31-mediated recombination between a construct containing the GFP/FlASH/StrepII/3×FLAG protein trap and an artificial MiMIC exon in the dysc locus (dysc Mi02729), as described previously (Venken
et al., 2011). Transgenic fly lines containing GFP/FlASH/StrepII/3×FLAG insertions were generated by standard germline transformation (Rainbow Transgenics). Fly head mRNA was extracted using TRIzol (Life Technologies), and reverse transcribed using High Capacity cDNA Reverse Transcriptase Kit (Applied Biosystems). See methods in the supplementary material for primer sequences used in this study. Larval behavior
Third-instar wandering larvae were collected, gently rinsed in PBS and placed on 5% sucrose/2% agar plates. One larva was counted per plate, and the number of peristaltic body wall contractions was measured visually over a 1 min period under a dissecting microscope. Confocal microscopy, immunohistochemistry and quantification of synaptic parameters
Wandering third instar larvae were used for confocal and STED imaging of larval synapses and AZs at muscle 6/7 or muscle 4 of segment 3. For all larval experiments, both sexes were used. Adult brains were dissected and immunostained as described previously (Wu and Luo, 2006). All confocal microscopy was performed using an Olympus Fluoview confocal microscope. Images were contrast enhanced in Adobe Photoshop. Where images of multiple genotypes are presented, all images were subject to the same enhancements. See methods in the supplementary material for further information on quantification of synaptic parameters, antibodies and staining protocols. STED imaging
STED images were acquired on a Leica TCS SP5 STED CW microscope with a 100×1.4 NA oil objective. The Alexa 488 secondary antibody was excited with a 488 nm laser line and depleted by a 592 nm laser line. STED images were deconvolved using the built-in deconvolution algorithm of the Leica LAS-AF software (Signal Intensity filter, regulation parameter: 0.010.02). The point spread function was generated using a 2D Lorentz function with the full-width half-maximum set to 70 nm. AZ areas were measured blind with respect to the experimental genotype. Only individual AZs that could be clearly distinguished from other AZs were included in the area measurements. Electrophysiology
Larvae were dissected and recorded in hemolymph-like saline HL3.1 containing (in mM): 70 NaCl, 5 KCl, 1 CaCl2, 4 MgCl2, 10 NaHCO3, 5 trehalose, 115 sucrose and 5 HEPES ( pH 7.2) as described previously (Feng et al., 2004; Ueda and Wu, 2006). The segmental nerves were severed from the ventral ganglion and stimulated with a suction electrode. Recordings were performed on ventral longitudinal muscle 6/7 in abdominal segments A3-A5 of third instar larvae. All cells selected for recording had resting membrane potentials between −50 and −70 mV. Both spontaneous and evoked postsynaptic currents were recorded while the muscle cell was voltage clamped at −60 mV, using an AxoClamp 2A (Axon Instruments) amplifier in single-electrode voltage clamp mode (switching frequency 10 KHz) and sharp microelectrodes (WPI) filled with 3 M KCl (5-10 MΩ resistance). Acknowledgements We thank Dr Nigel Atkinson and Bloomington and Harvard Drosophila stock centers for fly stocks; Dr Hugo Bellen and Dr Thomas Lloyd for technical advice; and Huihui Pan and Katelyn Kallas for technical assistance.
Competing interests The authors declare no competing financial interests.
Author contributions J.E.C.J., M.S., D.L., S.J.leM. and S.L. performed experiments. K.K., I.B.L., M.B.D. and M.N.W. supervised the work. J.E.C.J. and K.K. conceived the project and wrote the manuscript, with contributions from all authors.
Funding This work was supported by the National Institutes of Health [GM088221 (to K.K.); DA022727, MH08642 and MH100093 (to M.B.D.); NS17910 (to I.B.L.), NS079584
By what mechanisms might the DYSC/SLO complex influence AZ architecture? SLO channels contain a large intracellular Cterminal domain (Lee and Cui, 2010). Interestingly, recent studies have demonstrated that the C-terminal domain of SLACK channels, BK channel homologs that are potentiated by sodium rather than Ca2+, are bound to by the Fragile-X mental retardation protein in a manner that induces channel activation (Brown et al., 2010; Zhang et al., 2012). The C-terminal domain of SLO channels may similarly act as a scaffold to bind synaptic proteins that regulate AZ structure. Alternatively, DYSC may act as the key interacting node via its PDZ domains in a SLO-dependent manner. Distinguishing between these (and other) possibilities will be a productive avenue of future research. The morphological and physiological phenotypes observed in dysc and slo mutants may be mechanistically linked. Recent evidence suggests a causal link between spontaneous neurotransmission and synaptic bouton budding (Choi et al., 2014). Under genetic conditions where spontaneous neurotransmission is inhibited but evoked neurotransmission is maintained, hyper-budding of synaptic boutons is observed at the larval NMJ (Choi et al., 2014). Importantly, mutations in complexin, which result in extremely high frequencies of miniature events due to a loss of pre-fusion clamping of synaptic vesicles, exhibit increased synaptic bouton size (Choi et al., 2014; Huntwork and Littleton, 2007). The enhancement of spontaneous neurotransmission in dysc and slo mutants may therefore contribute to the enlargement of synaptic boutons in both mutant backgrounds. In addition, alterations in the synaptic localization of cytoskeletal protein Futsch may also act to independently increase synaptic bouton size in dysc and slo mutants. Futsch is a microtubule-binding protein that is crucial for synaptic bouton budding (Roos et al., 2000); mutations resulting in loss of Futsch function or the mislocalization of Futsch inhibit correct synaptic bouton budding, resulting in synapses with a reduced number of aberrantly large boutons (Mosca et al., 2012; Pennetta et al., 2002; Roos et al., 2000). We therefore posit a causal role for Futsch in mediating the abnormal development of synaptic boutons that results from loss of DYSC and SLO. However, we cannot rule out the possibility that mislocalization Futsch merely correlates with increases in bouton size in dysc and slo mutants. Mutations in futsch do not affect AZ structure or increase spontaneous neurotransmission (Lepicard et al., 2014), suggesting the alterations in T-bar architecture and spontaneous vesicle release are not downstream of changes in synaptic Futsch localization, but represent parallel pathways by which DYSC and SLO influence synaptic development and physiology. In conclusion, our research sheds new light on the roles of DYSC and its binding partner SLO in the Drosophila nervous system. The mammalian DYSC and SLO homologs whirlin and KCNMA1 may play similar roles in synaptic development and function.
Development (2014) 141, 1-10 doi:10.1242/dev.109538
(to M.N.W.)], by a grant from the Farber Family Foundation (I.B.L.) and by a Burroughs-Wellcome Fund Career Award for Medical Scientists (M.N.W.). Deposited in PMC for release after 12 months.
Supplementary material Supplementary material available online at http://dev.biologists.org/lookup/suppl/doi:10.1242/dev.109538/-/DC1
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RED FIFE WHEAT
What's so special about Red Fife?
Red Fife is quickly gaining popularity. It is characterized by an exceptional robust flavour and baking properties. Artisan bakers, pasta makers, health food stores and discerning food-origin conscious cooks who want to work with and use non-GMO products are searching the countryside for this unique heritage grain.
HISTORY OF RED FIFE
Red Fife wheat fed Canada for 60 years in the 1800's. By 1860 it was distributed across the country, adapting to a wide range of diverse growing conditions. Red Fife is the parent to several offspring wheat varieties, with "Marquis" being the #1 wheat in the early 1900's. Marquis, along with other derivatives ripened a few days earlier and eventually became the farmers choice for planting. Many of the bread wheat in Canada owe part of their genetic lineage to Red Fife and Canadian wheat are some of the finest bread wheat in the world.
Red Fife is a 'landrace' which means the kernels have uniform shape but with the genetic diversity, common to old varieties, able to adapt to diverse growing conditions. Often referred to as 'folk seeds' or 'farmer varieties', landraces tend to provide farmers with some yield regardless of seasonal conditions.
One legend states that a load of bread wheat originating in the Ukraine was on a ship in the Glasgow harbour. A friend of Ontario farmer David A. Fife accidentally dropped his hat into the red coloured wheat, collecting a few seeds in the hatband, which he then shipped off to the farmer Fife. The family cow managed to eat all the wheat heads except for one, which was the beginning of 'Red Fife' wheat in Canada.
RED FIFE REBORN
From 1900 to 1988 Red Fife was grown in very small quantities by plant breeders' seed collections. Then the 'Heritage Wheat Project' which began in 1988 was an initiative to revisit the heritage wheat varieties.
It is challenging to grow and there are very limited quantities being produced for the surging demand of Red Fife.
Learn more at these resources:
"Canada’s oldest successfully grown variety of wheat,
known as Red Fife, has found a home at the Ukrainian
Cultural Heritage Village..."
"Red Fife is a variety of bread wheat (Triticum aestivum) that David Fife and family began to grow in 1842 in Peterborough Ontario. Legend goes that a friend of Mr. Fife collected a sample..."
"Red Fife wheat is characterized by 3 little awns at the top of the wheat head but is otherwise awnless. The straws can be from 0.9 to 1.5 metres tall..."


frequently asked questions
Rivers and estuaries play an important role in delivering freshwater and materials such as sediments and contaminants to the coastal ocean. The role of waves in mixing and transporting this material along the coastline is not yet well understood. The non-toxic dye released in this experiment acts as a tracer for these freshwater outflows (plumes), which will help scientists better understand the fate of freshwater and land-derived materials from small plumes as they enter the coastal ocean and the surfzone. The improved understanding of this mixing in the coastal ocean is crucial to quantifying the spreading of sediment, pollutants, larvae, harmful algal blooms, and other important material near the coast where many people live and recreate.
The experiment will use an environmentally safe passive tracer called Rhodamine WT. The WT stands for “water tracer” because it is a water-soluble dye. The bright pink fluorescent dye occupies a very different range of wavelengths on the visible light spectrum than the background ocean water, making it straightforward to detect both with our eyes, with underwater sensors, and from above with optical cameras. In the water, an instrument called a fluorometer shines a green light to detect the red light emitted from the dye. The amount of red light is directly proportional to the concentration of the dye in the water, which scientists have calibrated with lab samples.
This chemical is not considered hazardous by the 2012 OSHA Hazard Communication Standard (29 CFR 1910.1200). The product contains no substances which, at their given concentration, are considered to be hazardous to health. This product is approved for use by the U.S Environmental Protection Agency (EPA) and Food and Drug Administration (FDA) in water tracing studies.
Scientists will release 15 gallons of dye during each of these dye releases. It will be visible to the naked eye in the water for several hours after the deployment, with small traces detectable by instruments for approximately 24 hours.
The project site is the estuary and surrounding coastline at Los Peñasquitos Lagoon, located within Torrey Pines State Beach and Natural Reserve in San Diego, California. The dye will be released near the estuary mouth during an ebb tide, the tidal phase when the water level is falling, so that it is carried out of the estuary into the coastal ocean. This site was chosen as a prime example of a small river plume discharging into the surfzone along a relatively uniform stretch of shoreline.
Yes, permits have been secured and approved from all necessary agencies.
This site was chosen because Los Peñasquitos Lagoon/Torrey Pines State Beach provides a prime example of a small river plume discharging into the surfzone along a relatively uniform stretch of coastline. The geography, wave, tide, weather, and river flow conditions make this location ideal for an experiment and a great example of these types of systems, which are found worldwide. Moreover, the location has been the site of many studies by other scientists such that the estuary and beach dynamics are relatively well understood. This site also benefits from a suite of long-term monitoring stations collecting complementary data (from USGS, NOAA, CDIP, NERR). This location was chosen for these physical properties and proximity to other measurements only.
The dye dispersal will be tracked in a variety of ways. Since the dye is fluorescent, scientists are able to track it using fluorometers that can measure the fluorescence or light emitted from the dye. These fluorometers, along with Conductivity-Temperature-Depth (CTD) sensors, which measure salinity and temperature, will be affixed to poles in the sand in the river mouth and surfzone. A jet ski outfitted with fluorometers, GPS and CTDs will also perform transects, or specified survey routes, along the shoreline. Offshore, beyond the breaking waves, several moorings have been deployed on the seafloor with CTDs, fluorometers, and instruments to measure ocean currents. Additionally, unpiloted aircraft systems (drones) will capture the structure and spatial extent of the dye plume from above, using RGB, hyperspectral, and infrared cameras. Beyond tracking of the dye, oceanographic conditions (waves, currents, tides, salinity, temperature, dissolved oxygen, and pH) will be monitored using sensors affixed to moorings deployed for this project.
The dye is nontoxic, so there are no harmful effects from being in the water. There will, however, be a lot of activity from researchers on the day of the dye release. Given this it may be advisable to recreate in an area further south or north of the estuary.
Scripps Institution of Oceanography scientists have successfully conducted nearshore dye experiments (using this same dye) at Huntington State Beach, Calif. (2006), Imperial Beach, Calif., (2009, 2015) and New River, N.C. (2013), and numerous experiments with surfzone, estuary, and offshore instruments. The City of San Diego also did a similar experiment in Mission Bay in 2021, where the dye was released from Rose Creek.
This experiment is sponsored by the National Science Foundation (Award #1924005 ). Complementary data at the same site examining estuarine dynamics and estuary mouth sedimentation is funded by the California Department of Parks and Recreation Natural Resources Division, Oceanography Program.
This project is led by scientists at UC San Diego’s Scripps Institution of Oceanography and the University of Washington. Additional collaborators include scientists from Universidad Técnica Federico Santa María (Chile), UC Irvine, UC San Diego Mechanical Engineering, and the Naval Postgraduate School. The lead investigator is coastal physical oceanographer Sarah Giddings.


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Why deep-sea exploration remains a great mystery
Since the dawn of humanity, we have always been fascinated by the unknown. While we have made remarkable progress in exploring the vastness of space, much of our own planet still remains a mystery. One of the most enigmatic and captivating areas that continue to intrigue scientists and adventurers alike is the deep-sea. In this article, we will delve into the complexities and uncertainties surrounding deep-sea exploration, shedding light on why this realm holds such allure.
The Depths of the Abyss: Unraveling the Unknown
The deep-sea, also known as the abyssal zone, refers to the vast and virtually unexplored area of the ocean lying beyond the reach of sunlight. As we descend into the depths, the pressure rapidly intensifies, creating an environment that is hostile to most forms of life. The darkness is impenetrable, and the extreme conditions pose numerous challenges for researchers. Despite advances in technology, our understanding of this mysterious realm remains limited.
One of the main reasons for this lack of knowledge is the immense depth and vastness of the ocean. With over 70% of our planet covered by water, exploring the deep-sea poses logistical challenges that are not easily overcome. The pressure at the ocean floor is crushing, reaching up to 1000 times greater than at the surface, which makes it difficult to develop robust equipment that can withstand such extreme conditions.
Moreover, the deep-sea is teeming with life forms that have adapted to survive in this hostile environment. From bioluminescent creatures to organisms that thrive near hydrothermal vents, the deep-sea is a haven of species that have evolved unique adaptations. Unraveling the complexities of these ecosystems and understanding their interconnectivity is a daunting task, requiring extensive research and collaborative efforts.
Exploring the Abyss: New Technologies and Discoveries
With each passing year, advancements in technology bring us closer to unraveling the secrets of the deep-sea. Remotely Operated Vehicles (ROVs) equipped with high-definition cameras and sophisticated sampling tools allow scientists to explore depths that were once deemed inaccessible. The use of robotic exploration vehicles has revolutionized our ability to study the deep-sea, providing unprecedented insights into its wonders.
Recent discoveries have astonished scientists and challenged our understanding of life on Earth. From the strange and otherworldly creatures found in the Mariana Trench, the deepest part of the ocean, to the discovery of undersea mountains and hidden ecosystems, each exploration mission brings new revelations. With every dive, we uncover more questions than answers, pushing the boundaries of our knowledge.
Despite the remarkable progress made in deep-sea exploration, much of the abyss remains uncharted. The sheer vastness of the ocean, combined with the challenges posed by extreme conditions and limited resources, means that there is still much to discover. The deep-sea holds the potential for groundbreaking scientific discoveries and the unlocking of mysteries that have eluded us for centuries.
The Final Frontier: Deep-Sea Exploration for a Sustainable Future
Deep-sea exploration not only satisfies our innate curiosity but also holds significant value for our future. The resources found in the deep-sea, such as rare metals and hydrocarbons, have the potential to fuel technological advancements and serve as a vital source for economic growth. Additionally, understanding the complex oceanic ecosystems is crucial for conservation efforts and the sustainable management of resources.
However, it is essential to strike a balance between exploration and preservation. The deep-sea is an incredibly delicate and fragile ecosystem, and any disturbances caused by human activities can have long-lasting effects. Responsible and ethical exploration, guided by scientific knowledge and environmental considerations, is paramount to ensure the preservation of this unique realm and the species that call it home.
In conclusion, deep-sea exploration continues to captivate our imagination, offering a glimpse into the mysterious and unknown. The challenges posed by extreme conditions and limited resources make the exploration of the abyss an ongoing endeavor, filled with uncertainty and excitement. As we dive deeper into the depths of the ocean, our thirst for knowledge grows, and the potential for groundbreaking discoveries beckons us forward. The deep-sea remains a great mystery, yet to be fully unraveled.


HUMAN ENDOCANNABINOID SYSTEM
Research suggests that regular doses of naturally occurring CBD may help relieve pain, speed recovery, reduce stress and regulate sleep.
Having only been discovered in recent decades, the endocannabinoid system (ECS) is a biological system composed of endocannabinoids (endo meaning they occur naturally in the body), or neurotransmitters, that bind to cannabinoid receptors present throughout the body. The main function of the ECS is to maintain bodily homeostasis, a state of internal balance and physical wellbeing.
Research has shown that the ECS is involved in a wide variety of processes including pain, memory, mood, appetite, stress, sleep, metabolism, immune and reproductive functions. Science has proven that endocannabinoids support the body’s inflammatory response by targeting CB1 receptors in the central nervous system and CB2 receptors found in the peripheral nervous systems, especially immune cells.
It has also been found that natural cannabinoids from Hemp and other plants help support the ECS and enhance its signaling. CBD is believed to augment the action of endocannabinoids thus supporting their effect on the body. This suggests that regular doses of naturally occurring cannabinoids might act as a tonic to our most central healing system.
Common Hemp CBD Health Benefits
Muscle and Joint Support
CBD has been found to modulate the levels of cytokines and chemokines—key mediators of the inflammatory response. In addition, CBD and other phytocannabinoids interact with vanilloid type-1 receptors (TRPV1), also called the capsaicin receptor, which is responsible for sensations of pain, heat and inflammation.
It is well-known that high stress can lead to problems with relaxation, sleep induction and sleep maintenance which all impact how easy it is to fall and stay asleep. In recent clinical evaluations, a majority of individuals with self-reported anxiety and poor sleep were found to have improved sleep scores after consuming CBD.
Anxiety and Stress Relief
Emerging evidence continues to explain the anxiolytic (anxiety reducing) effects of CBD, even though all of the details are yet to be defined. Preliminary research demonstrates that elevated doses of CBD can impact cortisol levels—which, when properly moderated, can help you deal more calmly with stressful situations.
Better your health | Better your adventures
Hemp CBD for Athletic Performance
We push ourselves both physically and mentally in sport, training and strenuous activity, which can lead to exhaustion, soreness and even pain. As an inflammation modulator, Hemp CBD has the potential to hasten muscle recovery and can help provide relief from occasional aches and pains which limit athletic performance.
Another essential component to physical fitness is sleep—it aids in both muscle and energy recovery. Hemp CBD has the potential to support sleep regularity by modulating cortisol levels that can heighten stress when not properly balanced. In turn, more restful sleep further strengthens the body’s response to stress in a type of CBD virtuous cycle.
Hemp CBD + Botanicals
While Hemp CBD has many physical and mental benefits—and can provide immense amounts of support—it’s not a cure-all for all ailments. At Windward, we combine Hemp with other therapeutic botanicals and adaptogens, which are plants that help the mind and body cope with stress. Our proprietary formulations create a greater effect than CBD alone, and we believe the synergy between plants and their active compounds is what truly enhances the entourage effect when taken in combination.
Learn More in our Botanical Glossary


Did you know that the Harrison River has one of the largest concentrations of bald eagles in North America in a one square mile radius? Yes they are right here in your backyard. With over 2000 eagles feeding on the spawning salmon in the area it truly is a sight to see. The best way to experience the eagles is by boat and two tour companies based in Harrison Hot Springs will take visitors to view the eagles and most likely you will also see heron, seals, bears and spawning salmon.
The Fraser Valley Bald Eagle Festival celebrates the bald eagle with their annual event Nov 21/22. Several sites have been established in and around the Eastern Fraser Valley with activities and viewing sites. Be sure to check them out at www.fraservalleybaldeaglefestival.ca. Conservationist and author David Hancock through the Hancock Wildlife Foundation have set several eagle cams in the area. Visit www.hancockwildlife.org to view them.


InnoVista Sensors™ selected biodiversity and bees as their first topic to launch the Environment Days in Valence. Wilfried Sanchez from the Rovaltain Foundation and Gabriel Faysse from Apiculture Urbaine presented the importance of bees, their role in food chain and their contribution on our world. We learnt that, each year, bees directly impact 153 milliards € in the agriculture market with 35% of our food tonnage strongly or totally dependant on animal pollination (from cultures to vegetables, fruit trees, oilseed crops,…). Meanwhile, the bee mortality rate in France is about 15% but reaches a frightening 80% in some parts of the world. Several factors are responsible for the disappearance of bees: pesticides are not the only ones to be blamed but also air pollution, multiplication of electromagnetic waves (high voltage powerlines for instance), as well as climatic changes, endocrine disruptors, etc. At the same time, apiculture has become global with swarms of bees being imported to Europe without paying attention to local seasonality nor to geographical specificity. This can directly impacts the way bees feed, their capacity to resist diseases, their fecundation and pollination rates. Gabriel Faysse, the beekeeper looking after our InnoVista Sensors™ beehives explains that they are hosted in the “Drôme des Collines” where Acacia trees, Lime trees, Chestnut trees and Lavender fields that are located within 3kms radius. Due to the very humid climatic conditions, our first honey jars will mostly be from chestnut and lavender flowers. We should receive our first delivery in September… Many thanks to Gabriel Fraysse and Wilfried Sanchez and to InnoVista employees who attended the conference.


You’ve just pushed through a hard workout. You are sweaty, sore, tired and (hopefully) feeling good about doing something beneficial for your mind and body. What should you do now to recover? Is there anything that you can do that will allow you to bounce back more effectively and capitalise on the work you have been doing in the gym? The short answer is yes, definitely.
What we eat and drink around our workouts and throughout our day can dramatically affect recovery rates. There are many nutrients and nutrient groups that help boost recovery, but there are four that are particularly important. These are water, electrolytes, protein and carbohydrates.
I think we all know by now how vital water is. Given how much of it we use during exercise, especially in things like sweat, it is important that we replace the amount we lose both during and after a workout. It is recommended to replace the water lost during a workout, plus 25 to 50% to help with recovery (weight lost on grams during workout x 1.25 to 1.5 = required millilitres of water). By having enough water in the body, toxins and byproducts left over from exercising can be removed more efficiently and nutrients can be delivered to the areas that need them the most.
Alongside water, we tend to use and secrete minerals such as potassium, sodium and calcium at an accelerated rate when exercising. Just like water, if levels of these minerals, or as they are more commonly known – electrolytes – fall too low, cognitive and physical function can be affected. This can lead to cramping, muscle soreness and in some rare cases, seizures. All of this will negatively impact recovery and health. So, after a long, sweaty session at the gym, or a run outside, it could be a good move to consume an electrolyte mix/ premade drink, or foods such as bananas, oranges and peanuts which are high in these much-needed minerals.
This is a big one for many people who work out. The focus should be on regularly consuming protein throughout the day to promote the building and repair of muscles, bones and other tissues. The amount needed per person per day depends on their sex, size and activity levels, but can range anywhere from 0.8g per kg of body weight through to as high as 2-2.4g per kg of body weight (even higher for some endurance athletes).
There is merit to consuming good quality protein shortly after a workout, as it helps replenish depleted levels and further encourages the building/repair process. Experts recommend anywhere from 20-40 grams of protein from sources such as poultry, Greek yoghurt, milk, nut butters, and plant and animal-based protein powders to name a few. Keep in mind we do absorb nutrients more effectively from whole foods compared to supplements, but protein powder can be more time-effective to consume, especially with a busy schedule.
For most moderate to high-intensity workouts, we predominantly use some form of carbohydrate to fuel our movements. Therefore, it is important to stock up before and after a workout to a. give us enough energy to perform the workout and b. recover and not feel so tired after the workout.
Simple carbohydrates can be great for a high, short release of energy, whereas complex carbohydrates are great for a longer-lasting, lower release of energy. Given how long it takes to digest complex carbohydrates, it is recommended to eat them two to three hours before a workout, mainly to avoid stomach cramps, whereas simple carbohydrates can be consumed 60 to 90 minutes before a workout as they are digested faster.
Good sources of both simple and complex carbohydrates include apples, bananas, blueberries, brown rice, potatoes, rolled oats and whole wheat pastas and breads.
Passive and active recovery
Of all the things spoken about in this article, sleep is probably the most important when it comes to recovery and repair after a workout. During sleep, our bodies clear out any leftover byproducts and toxins and dedicate more energy to supercharging the repair and building process. Without enough of it, or without the right type of it, our ability to bounce back after a hard workout and maximise our gains will be greatly diminished. As adults, we should aim for seven to nine hours of uninterrupted sleep every night, with children needing slightly more and older adults needing slightly less.
A great way to further improve recovery is to get a massage. Massage can help to alleviate sore and overworked muscles, increase range of motion and promote blood flow throughout the body, improving the delivery of nutrients. Plus, a massage promotes the release of endorphins, dopamine and serotonin, which help to reduce stress levels and promote feelings of well-being and relaxation.
Cold showers and ice baths
Some gymgoers and athletes regularly have cold showers and ice baths straight after a workout to promote recovery. It is thought that the constriction and dilation of blood vessels brought on by the different temperatures helps to remove waste products produced during a workout, which leads to less muscle soreness and a quicker recovery time. They have also been shown to stimulate the release of norepinephrine, which can provide a boost to mood and energy levels.
Cold showers and ice baths are not for everyone though and if you have any heart or cardiovascular condition/s, please speak to your healthcare provider before jumping into one.
Periodisation of training
Finally, as we become fitter and stronger, we should be able to lift more and handle more cardiovascular activity. But, if we are constantly pushing our bodies without adequate rest, our ability to recover and rebuild will diminish and the chances of injuring ourselves will increase. A good training program should include both periods of loading and de-loading to maximise recovery and gains.
As most muscle groups take 24 to 48 hours to recover after a workout, it is a good idea to rotate through targeting different muscle groups and training methods throughout the week, month and year. For most people, it is also good to have one to two days off from exercising every week to relax and recover. There is also nothing wrong with taking two to seven days off or working at a much lighter intensity every few months. This will further encourage recovery and often helps to break through training plateaus.
Hopefully, the information in this article helps you to maximise the results from your workout routine. Remember that what you eat and how you recover can be just as important for achieving your goals as what you are doing in the gym.
Bodybuilding.com. (n.d.) Protein Calculator: How Much Protein Do I Need? Retrieved September 10, 2023 from https://www.bodybuilding.com/fun/calpro.htm
Kummer, M. (2023, September 10). 11 Ice Bath Benefits (And Why Cold Plunging Works). Michael Kummer. https://michaelkummer.com/health/ice-bath-benefits/
Meyer, M., & Ball, J. (2023, January 7). The Best Carbs to Eat Before and After a Workout. Eating Well. https://www.eatingwell.com/article/8022615/best-carbs-to-eat-before-after-workout/#:~:text=Examples%20of%20complex%20carb-rich%20foods%20to%20load%20up,whole-wheat%20pasta%2C%20blueberries%2C%20raspberries%2C%20apples%2C%20potatoes%20and%20yams.
Quinn, E., Lau, M. (2023, May 9). 10 Tips to Speed Recovery After Exercise. A Guide To Recovering Post-Workout. Verywellfit. https://www.verywellfit.com/ways-to-speed-recovery-after-exercise-3120085
Sports Dieticians Australia. (n.d.). Fluids in Sport. Retrieved September 10, 2023, from https://www.sportsdietitians.com.au/factsheets/fuelling-recovery/fluids-in-sport/


"In general, because of poor survivability of these coronaviruses on surfaces, there is likely very low risk of spread from products or packaging that are shipped over a period of days or weeks at ambient temperatures. Coronaviruses are generally thought to be spread most often by respiratory droplets. Currently there is no evidence to support transmission of Covid-19 associated with imported goods and there have not been any cases of Covid-19 in the United States associated with imported goods," the CDC wrote on its website. NSW Health agrees. "The risk of this virus being present on imported packages or products is negligible," its website said.


ChE Assistant Professor Brian Chaplin was awarded a new NSF-grant entitled “Multifunctional Electro-reactive Membranes for Efficient Biomass Separation”. The grant is for $160,000 with an effective start date of 9/1/2016.
This collaborative research project will develop efficient, scalable, and multifunctional reactive electrochemical membranes (REMs) for sustainable biomass harvesting and production of value-added products derived from biomass (e.g., biofuels and specialty chemicals). Biomass separation using membrane filtration is characterized by high operational cost due to severe membrane fouling and the need for frequent backwashing. The proposed research will use the REMs to mitigate membrane fouling, pretreat biomass to facilitate economical harvesting, and remove the toxic inhibitors that will promote water reuse. The research outcomes will advance the fundamental science and engineering in biomass separation, and potentially lead to critical, transformative technologies for biomass and food processing, drinking water treatment, and biomolecule purification in the pharmaceutical industries.
This work will use algae as a model biomass organism, and will focus on characterizing membrane fouling, water reuse for algae regrowth, and electrochemical pre-treatment of algal cells. Specific research tasks include (1) synthesis and characterization of a suite of tailored monolithic or nanofibrous REMs for algal recovery; (2) evaluation of algae separation efficiency, permeate water treatment, and anti-fouling properties of REMs; (3) elucidation of algae cell disruption and underlying mechanisms of electrochemical oxidation using microfluidic experiments; and (4) experimental and modeling assessment of membrane fouling and regeneration.
The results of this project are anticipated to lead to transformative solutions that address the grand challenges at the energy-water nexus. First, the research will provide fundamental guidelines to the design of REMs with excellent filtration performance, flexible design, and durability of operation. Secondly, the research will promote water and nutrient reuse in waste streams or cultivation media, which reduces water or energy footprints of renewable energy production. Thirdly, the project will train and mentor at least three Ph.D. students and a large number of undergraduate and senior high school students that will be recruited from underrepresented groups in STEM at the three collaborating institutions.


The Effects of Sugar on the Body
Sugar, the sweet seductress that lingers on our palates and in our lives, is perhaps one of the most prevalent and yet misunderstood ingredients in our modern diets. Found in sugary snacks, decadent desserts, fizzy soft drinks, and hidden in the darkest corners of processed foods, sugar’s omnipresence is undeniable. But what if I told you that sugar’s effects on your body extend far beyond a fleeting indulgence of your sweet tooth? In this comprehensive exploration, we’re delving deep into the intricate world of sugar and its profound impacts on the human body. Brace yourselves, because this journey will take us through the sweet beginning, the tumultuous sugar rollercoaster, and the multi-faceted effects on the body that extend from the brain to your very cells.
The Sweet Beginning
Let’s start at the sweet beginning – the moment sugar touches your lips. As you savor that piece of cake or sip that sugary soda, your body embarks on a remarkable journey to break down the sweetness into something your cells can use for energy.
When you consume sugar, the body springs into action. Simple sugars like glucose and fructose are rapidly absorbed into the bloodstream, providing a quick surge of energy. This is why you might feel an almost instant pick-me-up after indulging in a sugary treat. However, the story doesn’t end here.
In response to rising blood sugar levels, your body activates a highly sophisticated system. The pancreas releases insulin, a hormone responsible for regulating blood sugar levels. Insulin acts as the key that unlocks your cells, allowing them to absorb glucose from the bloodstream. When everything works as intended, it’s a finely tuned process, and your blood sugar remains within a healthy range.
The Sugar Rollercoaster
But what happens when you consume excessive amounts of sugar? Here’s where the sugar rollercoaster comes into play.
Picture this: you enjoy a generous serving of your favorite sugary delight, and your blood sugar levels skyrocket. Suddenly, you’re full of energy and vitality. It’s the sugar rush. However, it’s short-lived. Soon enough, your blood sugar levels plummet, sending you on a sugar crash. You feel tired, sluggish, and often, you’re left craving even more sugar to replicate that initial high.
This cycle of spikes and crashes is what nutritionists and health experts refer to as the sugar rollercoaster, and it can wreak havoc on your body over time.
The Effects on the Body
Let’s delve into the brain’s response to sugar. Excessive sugar consumption can lead to addiction-like responses in this incredibly complex organ. When you consume sugar, it triggers the release of dopamine, often referred to as the “feel-good” neurotransmitter.
This surge of pleasure in response to sugar can be intense. It’s why sugary foods are so enticing. Over time, this pleasurable experience can lead to cravings and overconsumption, akin to the effects of certain addictive substances.
It’s not surprising that many people find it difficult to resist sugary treats when they stimulate the brain’s reward centers in such a powerful way. In fact, some researchers have even likened sugar addiction to drug addiction.
Your heart, the tirelessly pumping muscle that keeps you alive, is significantly affected by sugar consumption. High sugar intake is associated with an increased risk of heart disease. Here’s how it happens:
- Blood Pressure: Sugar, especially in the form of fructose, can contribute to high blood pressure. High blood pressure is a well-known risk factor for heart disease and stroke.
- Inflammation: Excessive sugar intake can lead to chronic inflammation in the body. Inflammation is a driving force behind many chronic diseases, including heart disease.
- Fatty Deposits: Over time, high sugar consumption can contribute to the buildup of fatty deposits in your arteries, a condition known as atherosclerosis. These fatty deposits can narrow and stiffen the arteries, making it harder for blood to flow and increasing the risk of heart attacks.
These factors combined make sugar a substantial contributor to heart disease, which remains one of the leading causes of death worldwide.
Your liver, a remarkable organ with a multitude of functions, plays a critical role in processing sugar. When you consume too much sugar, it can overload your liver and lead to a cascade of problems.
Fatty Liver Disease: One of the most common issues related to excessive sugar consumption is non-alcoholic fatty liver disease (NAFLD). As the name suggests, NAFLD causes the liver to accumulate excess fat, which can impair its function and even lead to more severe liver conditions.
Insulin Resistance: The constant demand placed on your liver to process sugar can also lead to insulin resistance. This is a condition where your cells become less responsive to insulin’s signals, causing your body to produce even more insulin to maintain blood sugar levels. Over time, this can lead to the development of type 2 diabetes.
Type 2 Diabetes: Long-term excessive sugar consumption can indeed lead to type 2 diabetes. In this condition, your body is unable to regulate blood sugar effectively, resulting in high and unstable blood sugar levels.
The battle against the bulge is one that many people face, and sugar can be a significant adversary. Sugar is calorie-dense, which means it’s packed with calories but devoid of essential nutrients. This makes it easy to consume more calories than you realize when indulging in sugary treats.
Moreover, sugary foods often lack the satiating power of whole, nutritious foods. They don’t fill you up as effectively, which can lead to overeating and, ultimately, weight gain. This is why you might find yourself reaching for that second or third cookie before feeling full.
The impact of sugar on weight gain goes beyond just its caloric content. The rollercoaster effect we discussed earlier, with its rapid spikes and crashes in blood sugar, can lead to increased hunger and cravings. This, in turn, makes it even more challenging to maintain a healthy weight.
The effects of sugar on dental health are a well-known fact, and you might have experienced it yourself. Sugar is like a feast for the harmful bacteria in your mouth. These bacteria thrive on sugar, producing acids that can erode your tooth enamel and lead to dental issues.
Over time, excessive sugar consumption can result in tooth decay, cavities, and gum disease. It’s not just about the quantity of sugar you consume; it’s also about the frequency of exposure. Constantly snacking on sugary foods or sipping on sugary drinks throughout the day exposes your teeth to repeated acid attacks, compounding the risk of dental problems.
Ways to Reduce Sugar Intake
Now that we’ve explored the extensive effects of sugar on the body, let’s talk about practical steps you can take to reduce your sugar consumption and make healthier choices. Remember, it’s not about complete deprivation, but rather finding a balanced approach that supports your health.
A great place to start is by becoming a savvy food label reader. Pay close attention to food labels and ingredient lists. Look for hidden sugars, which can go by various names such as high fructose corn syrup, sucrose, maltose, and any ingredient ending in “ose.”
Manufacturers often use different forms of sugar, making it challenging to spot added sugars. By scrutinizing food labels, you can make more informed choices and select products with less added sugar.
Choose Whole Foods
Opt for whole foods whenever possible. Fruits, vegetables, whole grains, and lean proteins should form the foundation of your diet. These natural foods contain naturally occurring sugars, along with essential nutrients and fiber.
The difference between the sugar in whole foods and added sugars is significant. Whole foods provide a slower, more balanced release of sugar into the bloodstream due to their fiber content. This helps regulate blood sugar levels and prevents the dramatic spikes and crashes associated with sugary processed foods.
Limit Sugary Beverages
Sugary beverages, such as soda, fruit juices, sports drinks, and even some seemingly innocent “health drinks,” are some of the worst culprits when it comes to sugar consumption. They can be a major source of added sugars in your diet.
Swap out sugary drinks for water, herbal teas, or other unsweetened beverages. This small change can significantly reduce your daily sugar intake while also benefiting your overall health.
While we’ve discussed the detrimental effects of excessive sugar consumption, it’s important to recognize that you don’t have to eliminate sugary treats from your life entirely. Enjoying the occasional dessert or sugary indulgence is a normal part of life.
The key is moderation. Be mindful of portion sizes and the frequency of your indulgences. Treat sugary foods as an occasional and special treat rather than a daily habit.
In conclusion, the effects of sugar on the body extend far beyond the temporary pleasure it provides to our taste buds. Excessive sugar consumption can lead to a host of health issues, including weight gain, heart disease, and even addiction-like responses in the brain.
It is crucial to be aware of the hidden sugars in our diets and to make informed choices to prioritize our health and well-being. Remember, moderation and balance are the keys to a healthier relationship with sugar. By adopting these practices, you can not only enjoy the occasional sweet treat but also protect your body from the adverse effects of excessive sugar consumption.
In a world where sugar is abundant and often hard to resist, knowledge and mindful choices become our greatest allies in the quest for better health. Your body will thank you for it in the long run. So, let’s embark on a journey towards a healthier, happier, and more balanced relationship with sugar – one that benefits both your body and your overall well-being.


Dec 19th 2007
The era of the hunter-gatherer was not the social and environmental Eden that some suggest
HUMAN beings have spent most of their time on the planet as hunter-gatherers. From at least 85,000 years ago to the birth of agriculture around 73,000 years later, they combined hunted meat with gathered veg. Some people, such as those on North Sentinel Island in the Andaman Sea, still do. The Sentinelese are the only hunter-gatherers who still resist contact with the outside world. Fine-looking specimens—strong, slim, fit, black and stark naked except for a small plant-fibre belt round the waist—they are the very model of the noble savage. Genetics suggests that indigenous Andaman islanders have been isolated since the very first expansion out of Africa more than 60,000 years ago.
About 12,000 years ago people embarked on an experiment called agriculture and some say that they, and their planet, have never recovered. Farming brought a population explosion, protein and vitamin deficiency, new diseases and deforestation. Human height actually shrank by nearly six inches after the first adoption of crops in the Near East. So was agriculture “the worst mistake in the history of the human race”, as Jared Diamond, evolutionary biologist and professor of geography at the University of California, Los Angeles, once called it?
Take a snapshot of the old world 15,000 years ago. Except for bits of Siberia, it was full of a new and clever kind of people who had originated in Africa and had colonised first their own continent, then Asia, Australia and Europe, and were on the brink of populating the Americas. They had spear throwers, boats, needles, adzes, nets. They painted pictures, decorated their bodies and believed in spirits. They traded foods, shells, raw materials and ideas. They sang songs, told stories and prepared herbal medicines.
They were “hunter-gatherers”.